- Email: [email protected]
Current views on eye development
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7 Colley, P.A. and Routtenberg, A. (1993) Brain Res. Rev. 18, 115–122 8 Alkon, D.L. (1995) Behav. Brain Res. 66, 151–160 9 Nogues, X., Micheau, J. and Jaffard, R. (1996) Behav. Brain Res. 75, 139–146 10 Wehner, J.M., Sleight, S. and Upchurch, M. (1990) Brain Res. 523, 181–187 11 Pascale, A. et al. (1994) Eur. J. Pharmacol. 265, 1–7 12 Fordyce, D.E. et al. (1995) Brain Res. 672, 170–176 13 Sunayashiki-Kusuzaki, K. et al. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 4286–4289 14 Van der Zee, E.A. et al. (1992) J. Neurosci. 12, 4808–4815 15 Dekker, L.V. and Parker, P.J. (1994) Trends Biochem. Sci. 19, 73–77 16 Newton, A.C. (1995) J. Biol. Chem. 270, 28495–28498 17 Nishizuka, Y. (1995) FASEB J. 9, 484–496 18 Abeliovich, A. et al. (1993) Cell 75, 1253–1262 19 Chen, C. et al. (1996) Cell 83, 1233–1242 20 Leitges, M. et al. (1996) Science 273, 788–791 21 Prekeris, R. et al. (1996) J. Cell Biol. 132, 77–90 22 Ishii, H. et al. (1996) Science 272, 728–731 23 Conrad, R. et al. (1994) J. Biol. Chem. 269, 32051–32054 24 Bell, R.M. and Burns, D.J. (1991) J. Biol. Chem. 266, 4661–4664 25 Gopalakrishna, R. et al. (1986) J. Biol. Chem. 261, 16438–16445 26 Wolf, M. and Sahyoun, N. (1986) J. Biol. Chem. 261, 13327–13332 27 Chapline, C. et al. (1993) J. Biol. Chem. 268, 6858–6861 28 Staudiger, J. et al. (1995) J. Cell Biol. 128, 263–271 29 Klauck, T.M. et al. (1996) Science 271, 1589–1592 30 Mochly-Rosen, D., Khaner, H. and Lopez, J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 3997–4000 31 Robles-Flores, M. and Garcia-Sainz, J.A. (1993) Biochem. J. 296, 467–472 32 Smith, B.L. and Mochly-Rosen, D. (1992) Biochem. Biophys. Res. Commun. 188, 1235–1240 33 Ron, D., Luo, J. and Mochly-Rosen, D. (1995) J. Biol. Chem. 270, 24180–24187 34 Kiley, S.C. and Jaken, S. (1994) Trends Cell Biol. 4, 223–227 35 Pascale, A. et al. (1996) J. Neurochem. 67, 2471–2477 36 Hashimoto, T. et al. (1988) J. Neurosci. 8, 1678–1683 37 Huang, F.L. et al. (1990) Dev. Brain Res. 52, 121–130 38 Hirata, M. et al. (1991) Dev. Brain Res. 62, 229–238 39 Tremblay, E., Ben-Ari, Y. and Roisin, M.P. (1995) J. Neurochem. 65, 863–870 40 Jiang, X. et al. (1994) Dev. Brain Res. 78, 291–295 41 Hunter, S.E., Seibenhener, M.L. and Wooten, M.W. (1995) Dev. Brain Res. 85, 239–248
42 Bothmer, J. and Jolles, J. (1994) Biochim. Biophys. Acta 1225, 111–124 43 Friedman, E. and Wang, H-Y. (1989) J. Neurochem. 52, 187–192 44 Meyer, M. et al. (1994) Neurobiol. Aging 15, 63–67 45 Battaini, F. et al. (1990) Neurobiol. Aging 11, 563–566 46 Battaini, F. et al. (1995) Neurobiol. Aging 16, 137–148 47 Fordyce, D.E. and Wehner, J.M. (1993) Neurobiol. Aging 14, 309–317 48 Narang, N. and Crews, F.T. (1995) Neurochem. Res. 20, 1119–1126 49 Pascale, A. et al. (1996) Neurosci. Lett. 214, 99–102 50 Battaini, F. et al. (1994) Biochem. Biophys. Res. Commun. 203, 1423–1431 51 Hrabetova, S. and Sacktor, T.C. (1996) J. Neurosci. 16, 5324–5333 52 Matsushima, H. et al. (1996) J. Neurochem. 67, 317–323 53 Battaini, F., Govoni, S. and Trabucchi, M. (1995) in Molecular Basis of Aging (Macieira-Coelho, A., ed.), pp. 269–291, CRC Press 54 Undie, A.S., Wang, H-Y. and Friedman, E. (1995) Neurobiol. Aging 16, 19–28 55 Pisano, M.R., Wang, H-Y. and Friedman, E. (1991) Biomed. Environ. Sci. 4, 173–181 56 Buchner, K. (1995) Eur. J. Biochem. 228, 211–221 57 Rogue, P.J., Ritz, M.F. and Malviya, A.N. (1993) FEBS Lett. 334, 351–354 58 Ron, D. et al. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 839–843 59 Gianotti, C. et al. (1993) Neurobiol. Aging 14, 401–406 60 Eckles, K.E. et al. (1995) Soc. Neurosci. Abstr. 21, 473 61 Mochly-Rosen, D. (1995) Science 268, 247–251 62 Jacken, S. (1994) in Protein Kinase C Current Concepts and Future Perspectives (Lester, D. and Epand, R.M., eds), pp. 237–254, Ellis Horwood 63 Faux, M.C. and Scott, J.D. (1996) Trends Biochem. Sci. 21, 312–315 64 Govoni, S. et al. (1992) Life Sci. 50, PL125–PL128 65 Lucchi, L. et al. (1993) Life Sci. 53, 1821–1832 66 Ron, D. and Mochly-Rosen, D. (1994) J. Biol. Chem. 269, 21395–21398 67 Krieger, C. et al. (1996) Trends Pharmacol. Sci. 17, 114–120 68 Manev, H. et al. (1990) FASEB J. 4, 2788–2797 69 Wang, H-J., Pisano, M.J. and Friedman, E. (1994) Neurobiol. Aging 15, 293–298 70 Masliah, E. et al. (1990) J. Neurosci. 10, 2113–2124 71 Nitsch, R.M. and Growdon, J.H. (1994) Biochem. Pharmacol. 47, 1275–1284 72 Govoni, S. et al. (1993) Neurology 43, 2581–2586 73 Bergamaschi, S. et al. (1995) Neurosci. Lett. 201, 1–4
Acknowledgements The authors wish to thank D. MochlyRosen for helpful comments, F.J. Van der Staay for providing the Wistar rats and W.C. Wetsel for the antisera to P.K.C. isoforms. This work was supported by the National Research Council (CNR) of Italy to F.B., the Ministry of University, Scientific and Technologic Research (MURST) to A.P., and by MURST and the Alzheimer Unit, Sacred Heart Hospital-FBF, Brescia, Italy to S.G.
Current views on eye development Guillermo Oliver and Peter Gruss Several genes involved in the regulation of eye development in different species have been identified. Structural and functional conservation have been found between some of these genes in organisms as diverse as Drosophila and mouse. One notable example is the relationship between the mouse Pax6 gene and eyeless of Drosophila. Ectopic expression of eyeless or mouse Pax6 in Drosophila results in the formation of additional eyes. Recently, another homeobox gene, Six3, was found to promote ectopic lens formation in fish embryos.The next step will be to unravel the associated regulatory pathways of these genes and assess the degree to which they display evolutionary conservation. This will be important in order to assimilate these findings with current anatomical and embryological models. It seems reasonable to believe that in the near future the characterization of the whole framework required for vertebrate eye development will be accomplished. Trends Neurosci. (1997) 20, 415–421
I
N SCIENCE AND MYTHOLOGY the eye has always been an exciting topic. Many open questions still remain, particularly those concerning the evolutionary aspects of how different organisms have solved the problem of forming and detecting an image. The identification of various conserved genes among organisms as different as Drosophila and mice has challenged some Copyright © 1997, Elsevier Science Ltd. All rights reserved. 0166 - 2236/97/$17.00
of the previously accepted ideas regarding the origin of metazoan visual system. How much of the development pathway leading to eye formation has been conserved during evolution remains to be answered. Here, we focus on some of the most recent aspects of metazoan eye development, with a particular emphasis on the role of certain homeobox genes. PII: S0166-2236(97)01082-5
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Guillermo Oliver is at the Dept of Genetics, St Jude Children’s Research Hospital, 332 North Lauderdale, Memphis, TN 38105, USA. Peter Gruss is at the Dept of Molecular Cell Biology, MaxPlanck Institute of Biophysical Chemistry, Am Fassberg, 37077 Göttingen, Germany.
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Fig. 1. Structure of the compound eye of Drosophila. (A) Scanning electron micrograph of a wild-type fly eye. (B) Schematic view of an ommatidial unit. A longitudinal section is shown on the left and cross sections at three different levels are shown on the right. Abbreviations: A, photoreceptor cell axons; AC, anterior cone cell; B, bristle; C, liquid-filled pseudocone; CZ, cone cells; EQC, equatorial cone cell; L, lens; M, basal membrane; PC, posterior cone cell; PLC, polar cone cell; PP, primary pigment cell; Rh, rhabdomere; SP, secondary pigment cells; TP, tertiary pigment cells; 1–8, photoreceptor cells R1–R8. Reproduced, with permission, from E. Hafen and K. Basler.
Evolutionary considerations An important, as yet unresolved aspect of visual system evolution is whether the similarities found in different eye structures arise as a consequence of a common ancestor or due to evolutionary convergence. In most living and fossilized animals it is possible to identify some kind of light-sensitive organ1. These structures vary from simple eye-spots with a small number of receptors to well-developed eyes2. The analyses of the various types of eyes in different animals indicate that for the majority of eye’s functional parts there have been alternative but similar solutions. Based on structure, anatomic origins and phylogeny, Salvini and Mayr1 proposed that the lens-containing eye of a vertebrate and the compound eye of an insect are independent evolutionary developments, and that eyes might have evolved independently as many as 40 different times. The main sensory component of all eye structures are the photoreceptor cells. These cells are classified as either ciliary or rhabdomeric, but no phylogenetic correlation can be made based on this criterion. However,
Fig. 2. Early development of optic cup and lens in vertebrate embryos. (A) Optic grooves start to evaginate in the diencephalon towards the surface ectoderm. (B) Optic vesicles contact the overlying ectoderm, which then differentiates into lens placodes. (C) The optic vesicle and the lens placode invaginate. They will become the neural and pigmented retina and the lens vesicle respectively.
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the fact that light absorption and phototransduction seem to be mediated by a common mechanism has supported the idea that photoreceptor cells have a monophyletic origin3. All visual systems, including those from organisms as diverse as insects, cephalopods and vertebrates, share similar light receptor molecules called opsins3,4. Opsins are seven-transmembrane receptors linked to a vitamin A-derived chromophore that is responsible for absorbing photons. This process is mediated by a G protein-linked signaling cascade in all living organisms. The conservation of opsins among different organisms suggests that photoreceptor cells have originated from a common ancestor and that a simple ancestral photosensitive cell evolved into the different types of eyes we see today4. Importantly however, G protein-linked receptors similar to opsins are known to function not only in light detection but also in many other different physiological processes5. Strong support for a common monophyletic origin has come from the discovery of some key regulator genes involved in the control of similar developmental pathways in Drosophila and vertebrate eye formation.
Eye formation and lens induction The eye structures of vertebrates and arthropods are perfect examples of distinct optical solutions to the problem often regarded as receiving and integrating an image. The compound eye of Drosophila and the eye and lens of vertebrates are clearly different, and both have been studied extensively. However, recent findings suggest that despite these apparent differences, the formation of the arthropod and vertebrate eyes rely on a conserved developmental pathway. Visual detection in adult Drosophila relies on the two compound eyes and the three dorsal ocelli. The compound eye of the adult fly (Fig. 1) consists of a hexagonal array of ~800 individual units or ommatidia, each containing eight photoreceptor neurons (R1–R8), 12 accessory cells, lens-secreting cone cells and a mechanosensory bristle4. Initially, the eye develops during the larval stage from the eye imaginal disc: an epithelial monolayer specified in the late embryo. Photoreceptor cells differentiate in the posterior margin of the eye disc and appear progressively in a wave-like fashion over a period of two days. The anterior movement of this wave of neural differentiation is called the ‘morphogenetic furrow’6. During the larval period, differentiation starts with the formation of the morphogenetic furrow, which progresses from posterior to anterior across the disc epithelium. Anterior to the furrow are the dividing, undifferentiated progenitor cells; immediately behind the furrow, cells form differentiating clusters; and more posteriorly, these clusters acquire their final differentiated state. The vertebrate eye (Fig. 2) is a very complex structure that originates from primordial tissues derived from the wall of the diencephalon and the overlying surface ectoderm. A functional lens ultimately requires the accumulation of lens crystallins7. Lens induction is often cited as a classic example of embryonic induction. One of the first experiments to document an inductive effect was performed by Spemann in 1901 (Ref. 8). Using a hot needle, he destroyed the optic vesicle primordium of the frog
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Rana temporaria and found that the optic anlage was required for lens formation. The result was that lenses did not develop, leading Spemann to conclude that the optic vesicle is necessary for lens formation. Spemann’s conclusions were later confirmed by Lewis9, who removed an optic vesicle from the head of a frog embryo at the late neurula stage and transplanted it below the flank ectoderm in areas that normally do not form lenses. He observed lens formation in association with the transplanted retinal tissue, and argued that the optic rudiment was sufficient for lens induction10. Subsequent experiments started to raise doubts about the generality of the so-called ‘optic-cup’ model of lens induction. By repeating Spemann’s experiments, King11 found that lenses could form in the absence of contact with optic vesicles. Moreover, other researchers found that in many species lenses can form if retinal tissue is removed at the neural plate stage (reviewed in Ref. 12). Furthermore, transplant experiments in which host and donor tissue was labeled showed that the optic vesicle was not sufficient for lens induction12–14. Many studies have been carried out in Xenopus to try to explain the mechanisms of lens induction. By tissue grafting and labeling, a current working model has been proposed (reviewed in Ref. 10). Based on this model, it is thought that in Xenopus the anterior neural plate acts as an early inducer of lens ectoderm during the late gastrula stage, and that lens induction involves four phases: competence; bias; specification; and differentiation. Competence is the capacity of a specific tissue, in this case the head ectoderm, to respond to an inducing signal. For example, in Xenopus, the animal cap ectoderm exhibits different competences according to the developmental stage (mesodermal, neural or lens). Transplantation experiments performed by Henry and Grainger in Xenopus10,13 showed that embryonic head ectoderm is competent to respond to lens-inductive interactions very early in development, during the mid-gastrula stage. They also showed that this process depends on the developmental timer and not on specific tissue interactions10. Later, Servetnick and Grainger15 showed that only mid-to-late gastrula ectoderm is competent to respond to signals coming from the anterior neural plate. At this stage, a planar signal coming from the anterior neural plate region might provide the initial lens determination signal10. Another inductive signal might also be provided by the region of the endoderm that underlies the lens area and that ultimately gives rise to the foregut. The combination of these signals thus enables the competent region of the head ectoderm to become committed or biased to a lens-forming fate10. In Xenopus, this bias is thought to be established at the neural plate stage and this condition increases as induction takes place. At some point during development, enough lens induction has occurred to bias the ectoderm to form a lens without further induction. As development proceeds, the optic vesicle contacts the presumptive lens region at the end of neurulation. The optic vesicle does not seem to be critical for the early lens-inductive events; however, it might be required to position the lens within the head ectoderm and during the later stages of lens differentiation10, as lenses formed without an optic vesicle are very rudimentary.
Do different eyes share common genes and pathways? The majority of the models of eye evolution and eye formation are based on morphological and embryological data. Recently, however, a variety of molecular markers expressed in different parts of the metazoan visual system have been identified and characterized. These markers have allowed us to decipher some of the key players in the complex process of eye formation, and have raised many new and important questions. Drosophila, Caenorhabditis elegans, mouse and zebra fish have become the models of choice for those trying to correlate embryological and molecular aspects of development. One of the advantages of these systems is the availability of numerous natural or induced developmental mutations, including some in the visual system. Molecular and genetic analyses of eye development in Drosophila have largely concentrated on the cellular interactions that regulate cell fate determination in the late larval eye disc16. A lot of data are available on the development of the different photoreceptors (R1–R8). Receptor tyrosine kinases, epidermal growthfactor-like molecules and Ras signal transduction are all required during photoreceptor development (reviewed in Ref. 17). Genes that regulate early events in fly eye development, within and anterior to the morphogenetic furrow have also been identified. In a region anterior to this furrow cells have been found to express patterning genes16. Some Drosophila eye mutants, such as eyeless, eyes absent, sine oculis and eye gone, are characterized by the partial or complete absence of compound eyes. These mutations are thought to affect patterning genes that act in the initial events of the regulatory cascade3. In addition, some secreted molecules are known to regulate pattern formation in the eye imaginal disc. For example, Decapentaplegic is required for proliferation and initiation of pattern formation at the posterior edge of the eye disc; it has also been suggested that it mediates hedgehog function during progression of the morphogenetic furrow across the disc18. In mice, a variety of eye mutants is available19 and new mutations have been generated by means of homologous recombination technology. Saturation mutagenesis in zebra fish has produced a collection of mutants affecting almost every aspect of embryonic development. This has given rise to some interesting new eye phenotypes, and the affected gene products are now awaiting isolation20. At the same time, many genes expressed during the development of the visual system have been isolated in other organisms. As a result, genes of developmental importance in flies have been found to have counterparts in vertebrates, and in some cases the reverse is also true (Table 1). These previously unsuspected relationships between Drosophila and vertebrates have been uncovered in various developmental pathways. One of the best-known examples is the conserved role of the Hox genes in the patterning of the antero–posterior body axis21. Many different groups of genes are expressed during different stages of eye development, including various homeobox genes (reviewed in Refs 22,23). Some homeobox-containing genes seem to have a conserved and important function during the development of TINS Vol. 20, No. 9, 1997
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TABLE 1. Genes involved in eye development Gene Organismsa (mouse/Drosophila) Pax6/eyeless Pax2/NF Otx2/orthodenticle Six3/sine oculis Chox10/NF Prox1/prospero
Gain-of-functionb
Loss-of-functionc
Ascidian, chicken, frog, fruitfly, human, Drosophila (ectopic eyes) mouse, nematodes, nemertean, quail, rat, Xenopus (axial defects) zebra fish Mouse, human, zebra fish
eyeless/reduced eyes (Drosophila) Small eye (mouse, rat) Aniridia (humans) Optic nerve coloboma (KO mouse) Optic nerve coloboma (human mutation) Chicken, frog, fruitfly, human, mouse, sea Xenopus (extra cement Defective head development and no 55 urchin, zebra fish gland) ocelli (Drosophila)53 Microphthalmia/cyclopia (KO mouse) Fruitfly, mouse Medaka fish (ectopic lens) Eyeless/reduced eyes (Drosophila) Goldfish, mouse, nematode Ocular retardation (mouse) Chicken, frog, fruitfly, human, mouse, Axonal outgrowth and GMC nematode defects (Drosophila)
Eya/eyes absent
Fruitfly, human, mouse, nematode62
NF/dachshund
Fruitfly
Drosophila (ectopic eyes)
Eyeless/reduced eyes (Drosophila)61 Branchio–oto–renal syndrome (mouse) Reduced or not eyes (Drosophila)43
Abbreviations: GMC, ganglion mother cell; KO, knockout; NF, not found. a Organisms in which homolog genes were found. b The results observed in those cases in which gain-of-function experiments were performed in different organisms. c The results observed either in natural occurring mutations (italics) or in those generated by in vitro inactivation of the gene. In both cases, special emphasis is only given to the visual system phenotype. References are only given for those cases not previously mentioned in the text.
the metazoan visual system; they are transcription factors that, aside from being highly conserved in evolution, play fundamental roles during early embryonic pattern formation21. The best-known example is represented by the Pax6/eyeless genes. Pax6 is a member of the Pax family of genes and contains two DNA-binding domains, a paired-domain and a homeodomain. Initially, this gene was isolated from a mouse genomic library by using other previously identified murine Pax genes as molecular probes24. Pax6 was found to be expressed early in development in the anterior neural plate, optic vesicles, lens and nasal placodes25,26. Later studies showed that, in mammals, mutations in this gene were responsible for the mouse Small eye phenotype and the human genetic syndrome aniridia27,28. Homozygous Small eye mutant embryos have several defects, such as being born without eyes and nose, and die soon after birth. This mutation represents one of the first correlations between a well characterized spontaneous mutation and a specific molecular marker. Interestingly, in the homozygous mouse embryos, the optic vesicles start to develop but then degenerate27,28. Early in mammalian development, Pax6 is expressed in the surface ectoderm that will form the lens placode. In Xenopus, Pax6 has been shown to be expressed at the late gastrula stage in the anterior border of the developing neural plate and in the prospective retinal tissue29. In homozygous Small eye mutants, whereas the optic vesicles start to form, the lens placode never develops; consequently, no eyes are present. Tissue recombination experiments showed that ectoderm from homozygous rSey/rSey (rSey is the rat counterpart of the Sey mouse) will not differentiate into lens tissue once cultured with normal optic vesicles30. By contrast, lens differentiation occurs with wild-type ectoderm and mutant optic vesicle30. This suggests that in mutant animals, the ability of the ectoderm to respond to an inductive signal is affected and that Pax6 participates in the estab418
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lishment of the competence in the head ectoderm. Therefore, it is possible that in Small eye mutants the optic vesicle degenerates because an essential signal, probably emanating from the lens placode area, is missing. Furthermore, apparently functional Pax6 binding sites have been found within the Pax6 promoter31 and in the promoters and enhancers of several crystallin genes22,32. Homologous Pax6 genes were isolated from several other species including sea urchin33, nemertean34, ascidian35, chicken36, quail37 and zebra fish38. Surprisingly, the well-known eyeless mutation in Drosophila was due to a mutation in a Pax6-related gene. This gene was isolated from a Drosophila expression library using an oligonucleotide corresponding to a homeodomain binding site as a probe39. The chromosomal localization of the Pax6 homolog indicated that it corresponded to the Drosophila eyeless mutation39. The eyeless gene is expressed in eye progenitor cells and during differentiation; it is strongly expressed anterior to and within the morphogenetic furrow39; and is also expressed during Drosophila nervous system development. Homozygous eyeless flies show a reduction or lack of eyes. These findings indicate that despite the clear anatomical differences between the compound Drosophila eye and the vertebrate eye, some conservation at the gene level has been maintained. Further support for this idea comes from the report of induction of ectopic eyes in Drosophila by targeted expression of either the fly eyeless gene product or the mouse Pax6 gene in the wings, legs and antennae imaginal discs40. However, misexpression of Pax6 during early Xenopus development resulted in axial defects, but no specific eye phenotype29. These results, together with the isolation of Pax6related genes in such evolutionary distant species as nemertines, ascidians, sea urchin and nematodes3, strongly suggest that Pax6/eyeless is one of the major
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players in visual system development in metazoans3,40. Interestingly, the nematode C. elegans, in which a Pax6 homolog gene has been identified41,42 has no eyes. In this case, Pax6 is involved in the patterning of the head region and it is also required for the development of a tail sense organ41,42. Targeted mis-expression has been performed in Drosophila with the product of the dachshund gene43. This gene encodes a nuclear protein required for normal eye formation in the fly. Similar to the result obtained with eyeless, mis-expression of dachshund induced ectopic eye formation in Drosophila43. Furthermore, the authors showed that dachshund and eyeless induce the expression of each other, and that dachshund is required for ectopic retinal development driven by eyeless mis-expression. No vertebrate counterpart for dachshund has yet been identified. Pax2, another member of the Pax family of transcription factors is also expressed during murine eye development44,45. Its expression is confined to those cells within the optic vesicle that contribute to the optic stalk. Pax2 knockout mice have coloboma, a failure of the optic fissure to close, indicating that this gene is required for closure of the fissure45. Human46 and zebra fish47 Pax2 homologs have also been identified. A Pax2 natural mutation in humans was found to be associated with abnormal choroid fissure formation46. Experiments with zebra fish homologs of these two Pax genes have shown that two members of the hedgehog family of signaling molecules, normally expressed in the midline, can promote Pax2 expression and inhibit Pax6 within the optic vesicle48,49. These data suggest that the hedgehog proteins, directly or indirectly, regulate Pax2 and Pax6 expression. Recent studies also indicate that a single morphogenetic field gives rise to the two separate retina primordia under the influence of the prechordal plate50. Sonic hedgehog, a gene expressed in the midline, seems a strong candidate for this retina patterning, probably by suppressing Pax6 expression in the medial part of the neural plate50. In Drosophila, the hedgehog gene is expressed in differentiating cluster cells but also influences events anterior to the morphogenetic furrow. Flies mutant for this gene show reduced eyes and cell death. Two vertebrate homologs of the Drosophila homeobox-containing gene orthodenticle have been isolated (Otx1 and Otx2) and both are expressed in the head and brain of the developing mouse embryo51. However, both genes are also expressed in the developing nasal epithelium and in different parts of the developing visual system51. Disruption of the Otx2 gene in mice causes microphthalmia, a phenotype associated with the eyes either protruding over the face surface or buried in the orbit. The retina is hyperplastic, and in some cases only one eye is present, usually without lens and cornea52. This suggests that Otx2 participates in some stages of eye development possibly by specifying anterior head formation. In Drosophila, orthodenticle is expressed in the most anterior part of the head, and it is also required for the formation of the ocelli in the adult fly53. Otx2-related genes were also identified in sea urchin, zebra fish, humans (see Ref. 54), Xenopus55 and chicken56. Chx10 is another murine homeobox gene expressed in the optic vesicles and neuroretina, particularly in
the inner nuclear layer57. Homologous genes were isolated in goldfish and C. elegans58. A naturally occurring mouse mutation in this gene was shown to be responsible for the ocular retardation phenotype (reduced cell proliferation rate in the neural retina)57. These data support the idea that Chx10 is essential for retinal cell proliferation and for bipolar cell and rod photoreceptor differentiation59. Three murine homologs of the Drosophila eyes absent gene have been isolated60; in flies, the eyes absent gene is expressed anterior to the furrow and encodes a novel nuclear protein required for eye development61; the murine homologs Eya1 and Eya2 are expressed during the development of the cranial placodes, and all three are expressed in the developing eye60. Mutations in the human Eya1 gene have been found to be responsible for the branchio–oto–renal syndrome62. Another interesting parallel between Drosophila and vertebrate eyes is represented by the prospero/Prox1 genes. The product of the fly prospero gene is expressed in the lens-secreting cone cells63, whereas its putative murine homolog, Prox1 is expressed in the lens fibers64. This suggests that lens development in different species shares some common conserved molecular mechanisms. Prospero-related genes have also been identified in C. elegans65, chicken66, human67 and Xenopus (G. Oliver, unpublished). The product of the Drosophila sine oculis gene (a homeobox-containing gene) is expressed anterior to the furrow and it is essential for the development of the entire visual system by controlling the initial events of pattern formation in the eye disc16. The sine oculis gene is expressed before patterning and its product is required for formation of the optic lobes. In some sine oculis mutants, ommatidia fail to develop, suggesting that this gene is essential for the initiation of pattern formation in the disc. These mutant flies also exhibit apoptosis anterior to the morphogenetic furrow and their optic lobe primordia do not invaginate16. Recently, a mammalian gene family with high sequence similarity to the sine oculis fly gene was identified68. This family contains at least five members, one of which, Six3, seems to be the murine functional homolog of sine oculis69. Interestingly, in mouse, the expression of some of the Six genes co-localizes in many tissues with that of Eya1 and Eya2 (Refs 60,68,69). Six3 is expressed in the anterior neural plate and prospective eye field (Fig. 3), optic stalk, optic vesicles, lens and nasal placode69. This expression pattern, together with the fact that the anterior neural plate is involved in Xenopus lens induction10, suggests that Six3 has a role in vertebrate eye formation. In an attempt to understand this role, we ectopically expressed mouse Six3 in fish embryos70. We found that Six3 was sufficient to promote ectopic lens formation in the area of the otic vesicle (Fig. 4) and that retinal tissue is not a prerequisite for ectopic lens differentiation. The finding that ectopic expression of a single mouse gene (Six3) promotes lens formation in a fish embryo demonstrates that this gene is a key player in this process70. The results also suggest that, like Pax6 in vertebrates and insects, Six3/sine oculis is also evolutionarily conserved in metazoan eye development. The lens placode is one of the sensory placodes that is represented by thickening of the head ectoderm surrounding the neural plate. In addition to the lens TINS Vol. 20, No. 9, 1997
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Fig. 3. (Left) Six3 early expression in the prospective eye field. Whole mount in situ hybridization of an 8.5-day-old mouse embryo (ventral view). Purple staining indicates Six3 expression in the anterior neural plate region. Eyes will later develop in that region. Fig. 4. (Right) Ectopic Six3 expression promotes lens formation in the otic vesicle territory. Three-day-old medaka embryo showing an ectopic lens (arrow) replacing the left otic vesicle (OV). The lens (L), lens epithelium (arrowhead) and neuroretina (NR) of the wild-type eyes are shown. Whole mount in situ hybridization analysis was performed on this embryo leading to an unspecific brown staining of the lenses.
placode, the other sensory placodes are: the nasal placode, which gives rise to the nasal epithelium; and the otic placode, which forms the inner ear. Jacobson proposed that initially a common equipotential placodal state is activated in a large region of head
ectoderm, and later, via different interactions with the developing brain, the specific sensory placodes form71. The induction of ectopic lenses in the otic placode territory demonstrates that the region of the otic placode is competent to form lenses in response to Six3, and supports previous tissue transplantation data72. Based on the proposed model for lens induction discussed above10, it is possible that ectopic Six3 expression changes the bias of the otic placode towards the lens pathway by inducing the secretion of some unknown factor70 (Fig. 5). In this case, an ectopic lens could form in the area in which the inner ear would normally develop. It is possible that during early development, the whole head ectoderm is competent to respond to a variety of different signals. The establishment of an unreversible commitment (bias) towards optic, otic or nasal placodes would thus depend on which signal or signals were available. In this scenario, Six3 could play a crucial role in the pathway of lens placode determination (Fig. 5). By analogy, one might predict that genes will be identified that can induce otic or nasal structures when expressed ectopically in uncommitted head ectoderm. To confirm the function of the Six3 gene in eye development we are currently working on the generation of Six3 null-mutant mice. Finally, it is still not yet clear whether the different metazoan eyes have evolved from a common ancestor or whether they have evolved independently on many different occasions using generic signaling pathways – in this case, similar molecules were recruited for this purpose. It is possible that some of the genes discussed previously (Pax6, Otx2, Six3 and Eya) were initially involved in the patterning of the anterior part of the embryo. The identification of a Pax6 homolog in an organism such as C. elegans, devoid of eyes, and its suggested role in the patterning of the anterior head structures also supports this proposal. Further on in evolution, the aforementioned genes become involved in the induction and differentiation of cranial placodes; in this respect, not only are they similarly expressed in anterior forebrain and eyes, but also in the nasal placodes and nasal epithelium, and some members of this group are also expressed in the otic placode and pituitary. Later, these and other still-unidentified genes might have been recruited to regulate different conserved structural molecules involved in light or other stimulus detection. From there, the different types of metazoan eyes might have evolved. It will be extremely interesting to find out whether the regulatory cascades in which these genes function have also been conserved during the evolution of eye development. Selected references
Fig. 5. Proposed model of ectopic lens induction by ectopic Six3 expression in fish embryos. According to the current model for lens induction in Xenopus10, in the wild-type situation, early in development the head ectoderm is still uncommitted. Later, and via different molecules, a competent phase for each of the sensory placodes is acquired. Finally, each of the placodes becomes unreversibly biased: nose, internal ears or lens will be specified and they will differentiate. In the case of the transgene, the ectopic overexpression of Six3 has probably affected the transition from competence to bias. As a result, what could eventually be considered a homeotic transformation occurred, and a lens was induced instead of the otic placode. Abbreviations: Fb, forebrain; Hb, hindbrain; Mb, midbrain.
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1 Salvini-Plawen, L.V. and Mayr, E. (1977) Evol. Biol. 10, 207–263 2 Land, M.F. and Fernald, R.D. (1992) Annu. Rev. Neurosci. 15, 1–29 3 Halder, G., Callaerts, P. and Gehring, W.J. (1995) Curr. Opin. Genet. Dev. 5, 602–609 4 Zuker, C.S. (1994) Science 265, 742–743 5 Nilsson, D.E. (1996) Curr. Biol. 6, 39–42 6 Wolff, T. and Ready, D.F. (1993) in The Development of Drosophila melanogaster (Bate, M. and Martinez-Arias, A., eds), pp. 1277–1326, Cold Spring Harbor Laboratory Press 7 Wistow, G. (1993) Trends Biochem. Sci. 18, 301–306 8 Spemann, H. (1901) Verb. Anat. Ges. 15, 61–79 9 Lewis, W. (1904) Am. J. Anat. 3, 505–536 10 Grainger, R.M. (1992) Trends Genet. 8, 349–355 11 King, H.D. (1905) Arch. Entw. Mech. 19, 85–107 12 Saha, M.S., Spann, C.L. and Grainger, R.M. (1989) Cell Diff. Dev. 28, 153–172 13 Henry, J. and Grainger, R. (1987) Dev. Biol. 124, 200–214
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14 Grainger, R., Henry, J. and Henderson, R. (1988) Development 102, 517–526 15 Servetnick, M. and Grainger, R. (1991) Development 112, 177–188 16 Cheyette, B.N.R. et al. (1994) Neuron 12, 977–996 17 Zipursky, L. and Rubin, G. (1994) Annu. Rev. Neurosci. 17, 373–397 18 Pignoni, F. and Zipursky, L. (1997) Development 124, 271–278 19 Graw, J. (1996) Dev. Genet. 18, 181–197 20 Malicki, J. et al. (1996) Development 123, 263–273 21 De Robertis, E.M., Oliver, G. and Wright, C.V.E. (1990) Sci. Am. 262, 46–52 22 Cvekl, A. and Piatigorsky, J. (1996) BioEssays 18, 621–630 23 Beebe, D.C. (1994) Invest. Ophthalmol. Vis. Sci. 35, 2897–2900 24 Walther, C. et al. (1991) Genomics 11, 424–434 25 Walther, C. and Gruss, P. (1991) Development 113, 1453–1449 26 Grindley, J.C. et al. (1995) Development 121, 1433–1442 27 Hill, R.E. et al. (1991) Nature 354, 522–525 28 Ton, C.C.T. et al. (1991) Cell 67, 1059–1074 29 Hirsch, N. and Harris, W.A. (1997) J. Neurobiol 32, 45–61 30 Fujiwara, M. et al. (1994) Differentiation 57, 31–38 31 Plaza, S. et al. (1995) Mol. Cell. Biol. 15, 892–903 32 Richardson, J., Cvekl, A. and Wistow, G. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 4676–4680 33 Czerny, T. and Busslinger, M. (1995) Mol. Cell. Biol. 15, 2858–2871 34 Loosli, F., Kmita-Cunisse, M. and Gehring, W.J. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 2657–2663 35 Glardon, S. et al. (1997) Development 124, 817–825 36 Goulding, M.D., Lumsden, A. and Gruss, P. (1993) Development 117, 1001–1016 37 Dozier, C. et al. (1993) Cell Growth Differ. 4, 281–289 38 Krauss, S. et al. (1991) EMBO J. 10, 3609–3619 39 Quiring, R. et al. (1994) Science 265, 785–789 40 Halder, G., Callaerts, P. and Gehring, W.J. (1995) Science 267, 1788–1792 41 Chisholm, A.D. and Horvitz, H.R. (1995) Nature 377, 52–55
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Zhang, Y. and Emmons, S.W. (1995) Nature 377, 55–59 Shen, W. and Mardon, G. (1997) Development 124, 45–52 Nornes, H.O. et al. (1990) Development 109, 797–809 Torres, M., Gomez-Pardo, E. and Gruss, P. (1996) Development 122, 3381–3391 Sanyanusin, P. et al. (1995) Nat. Genet. 9, 358–364 Krauss, S. et al. (1991) Development 113, 1193–1206 Ekker, S.C. et al. (1995) Curr. Biol. 5, 944–955 Macdonald, R. et al. (1995) Development 121, 3267–3278 Li, H. et al. (1997) Development 124, 3603–3615 Simeone, A. et al. (1993) EMBO J. 12, 2735–2747 Matsuo, I. et al. (1995) Genes Dev. 9, 2646–2658 Finkelstein, R. et al. (1990) Genes Dev. 4, 1516–1527 Finkelstein, R. and Boncinelli, E. (1994) Trends Genet. 10, 310–315 Blitz, I.L. and Cho, K.W.Y. (1995) Development 121, 993–1004 Bally-Cuif, L. et al. (1995) Mech. Dev. 49, 49–63 Liu, I.S. et al. (1994) Neuron 13, 377–393 Svendsen, P.C. and McGhee, J.D. (1995) Development 121, 1253–1262 Burmeister, M. et al. (1996) Nat. Genet. 12, 376–384 Xu, P-X. et al. (1997) Development 124, 219–231 Bonini, N.M. and Choi, K-W. (1995) Curr. Opin. Genet. Dev. 5, 507–515 Abdelhak, S. et al. (1997) Nat. Genet. 15, 157–164 Chu-Lagraff, Q. et al. (1991) Development (Suppl.) 79–85 Oliver, G. et al. (1993) Mech. Dev. 44, 3–16 Buerglin, T.R. (1994) Trends Biochem. Sci. 19, 70–71 Tomarev, S.I. et al. (1996) Dev. Dyn. 206, 354–367 Zinovieva, R.D. et al. (1996) Genomics 35, 517–522 Oliver, G. et al. (1995) Development 121, 693–705 Oliver, G. et al. (1995) Development 121, 4045–4055 Oliver, G. et al. (1996) Mech. Dev. 60, 233–239 Jacobson, A.G. (1966) Science 152, 25–34 Barbanov, V.M. and Fedtsova, N.G. (1982) Differentiation 21, 183–190
Acknowledgements We thank S. Vallance, T. Curran and L. Zipursky for critical reading of the manuscript, M. Kessel for helping in the preparation of Fig. 5, and an anonymous reviewer for some helpful suggestions.
Development of the insect stomatogastric nervous system Volker Hartenstein The stomatogastric nervous system (SNS) forms a network of peripheral ganglia associated with the insect gut.The SNS originates from a neuroepithelial placode which dissolves into a population of migrating neural precursors. The formation of the SNS presents many parallels to the development of the vertebrate peripheral nervous system. Recent studies have started to provide answers for pertinent questions in SNS development,in particular,how the SNS placode is specified, how SNS precursors are released in a reproducible pattern from this placode and how different cell types in the SNS are determined. Trends Neurosci. (1997) 20, 421–427
C
OMPARABLE IN FUNCTION to the autonomic nervous system of vertebrate organisms, there exists in invertebrates a system of peripheral ganglia and nerve fibres that innervate the visceral organs and control food digestion, respiration and excretion. In higher invertebrates, including annelids and arthropods, the anterior portion of the visceral nervous system surrounding the foregut consists of a chain of ganglia which are connected in a characteristic manner with the brain and endocrine system. These nerve centres innervate muscles of the mouth cavity, foregut and midgut, and regulate food uptake and food transport; they are referred to as the stomatogastric nervous system (SNS). Due to its small cell number, simple behavioral output and easy accessibility, the SNS of crustaceans has
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been used for many years as a model system to study the functioning of a neural network1. More recently, the development of the stomatogastric nervous system in insects has attracted the attention of several investigators2–9, in part due to the fact that in some aspects it resembles neural development in vertebrates. The vertebrate autonomic nervous system is derived from the neural crest. During neurulation, the neurectoderm (neural plate) invaginates and forms the neural fold. Most of the neural fold closes dorsally to form the neural tube (giving rise to the CNS); cells at the lateral fringes of the fold move out of the epithelium and become the neural crest10,11. Subsequently, these cells migrate along different routes and form the various ganglia of the sensory and autonomic nervous PII: S0166-2236(97)01066-7
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Volker Hartenstein is at the Dept of Molecular, Cell and Developmental Biology, University of California, Los Angeles, Los Angeles, CA 90095-1606, USA.
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7 Colley, P.A. and Routtenberg, A. (1993) Brain Res. Rev. 18, 115–122 8 Alkon, D.L. (1995) Behav. Brain Res. 66, 151–160 9 Nogues, X., Micheau, J. and Jaffard, R. (1996) Behav. Brain Res. 75, 139–146 10 Wehner, J.M., Sleight, S. and Upchurch, M. (1990) Brain Res. 523, 181–187 11 Pascale, A. et al. (1994) Eur. J. Pharmacol. 265, 1–7 12 Fordyce, D.E. et al. (1995) Brain Res. 672, 170–176 13 Sunayashiki-Kusuzaki, K. et al. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 4286–4289 14 Van der Zee, E.A. et al. (1992) J. Neurosci. 12, 4808–4815 15 Dekker, L.V. and Parker, P.J. (1994) Trends Biochem. Sci. 19, 73–77 16 Newton, A.C. (1995) J. Biol. Chem. 270, 28495–28498 17 Nishizuka, Y. (1995) FASEB J. 9, 484–496 18 Abeliovich, A. et al. (1993) Cell 75, 1253–1262 19 Chen, C. et al. (1996) Cell 83, 1233–1242 20 Leitges, M. et al. (1996) Science 273, 788–791 21 Prekeris, R. et al. (1996) J. Cell Biol. 132, 77–90 22 Ishii, H. et al. (1996) Science 272, 728–731 23 Conrad, R. et al. (1994) J. Biol. Chem. 269, 32051–32054 24 Bell, R.M. and Burns, D.J. (1991) J. Biol. Chem. 266, 4661–4664 25 Gopalakrishna, R. et al. (1986) J. Biol. Chem. 261, 16438–16445 26 Wolf, M. and Sahyoun, N. (1986) J. Biol. Chem. 261, 13327–13332 27 Chapline, C. et al. (1993) J. Biol. Chem. 268, 6858–6861 28 Staudiger, J. et al. (1995) J. Cell Biol. 128, 263–271 29 Klauck, T.M. et al. (1996) Science 271, 1589–1592 30 Mochly-Rosen, D., Khaner, H. and Lopez, J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 3997–4000 31 Robles-Flores, M. and Garcia-Sainz, J.A. (1993) Biochem. J. 296, 467–472 32 Smith, B.L. and Mochly-Rosen, D. (1992) Biochem. Biophys. Res. Commun. 188, 1235–1240 33 Ron, D., Luo, J. and Mochly-Rosen, D. (1995) J. Biol. Chem. 270, 24180–24187 34 Kiley, S.C. and Jaken, S. (1994) Trends Cell Biol. 4, 223–227 35 Pascale, A. et al. (1996) J. Neurochem. 67, 2471–2477 36 Hashimoto, T. et al. (1988) J. Neurosci. 8, 1678–1683 37 Huang, F.L. et al. (1990) Dev. Brain Res. 52, 121–130 38 Hirata, M. et al. (1991) Dev. Brain Res. 62, 229–238 39 Tremblay, E., Ben-Ari, Y. and Roisin, M.P. (1995) J. Neurochem. 65, 863–870 40 Jiang, X. et al. (1994) Dev. Brain Res. 78, 291–295 41 Hunter, S.E., Seibenhener, M.L. and Wooten, M.W. (1995) Dev. Brain Res. 85, 239–248
42 Bothmer, J. and Jolles, J. (1994) Biochim. Biophys. Acta 1225, 111–124 43 Friedman, E. and Wang, H-Y. (1989) J. Neurochem. 52, 187–192 44 Meyer, M. et al. (1994) Neurobiol. Aging 15, 63–67 45 Battaini, F. et al. (1990) Neurobiol. Aging 11, 563–566 46 Battaini, F. et al. (1995) Neurobiol. Aging 16, 137–148 47 Fordyce, D.E. and Wehner, J.M. (1993) Neurobiol. Aging 14, 309–317 48 Narang, N. and Crews, F.T. (1995) Neurochem. Res. 20, 1119–1126 49 Pascale, A. et al. (1996) Neurosci. Lett. 214, 99–102 50 Battaini, F. et al. (1994) Biochem. Biophys. Res. Commun. 203, 1423–1431 51 Hrabetova, S. and Sacktor, T.C. (1996) J. Neurosci. 16, 5324–5333 52 Matsushima, H. et al. (1996) J. Neurochem. 67, 317–323 53 Battaini, F., Govoni, S. and Trabucchi, M. (1995) in Molecular Basis of Aging (Macieira-Coelho, A., ed.), pp. 269–291, CRC Press 54 Undie, A.S., Wang, H-Y. and Friedman, E. (1995) Neurobiol. Aging 16, 19–28 55 Pisano, M.R., Wang, H-Y. and Friedman, E. (1991) Biomed. Environ. Sci. 4, 173–181 56 Buchner, K. (1995) Eur. J. Biochem. 228, 211–221 57 Rogue, P.J., Ritz, M.F. and Malviya, A.N. (1993) FEBS Lett. 334, 351–354 58 Ron, D. et al. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 839–843 59 Gianotti, C. et al. (1993) Neurobiol. Aging 14, 401–406 60 Eckles, K.E. et al. (1995) Soc. Neurosci. Abstr. 21, 473 61 Mochly-Rosen, D. (1995) Science 268, 247–251 62 Jacken, S. (1994) in Protein Kinase C Current Concepts and Future Perspectives (Lester, D. and Epand, R.M., eds), pp. 237–254, Ellis Horwood 63 Faux, M.C. and Scott, J.D. (1996) Trends Biochem. Sci. 21, 312–315 64 Govoni, S. et al. (1992) Life Sci. 50, PL125–PL128 65 Lucchi, L. et al. (1993) Life Sci. 53, 1821–1832 66 Ron, D. and Mochly-Rosen, D. (1994) J. Biol. Chem. 269, 21395–21398 67 Krieger, C. et al. (1996) Trends Pharmacol. Sci. 17, 114–120 68 Manev, H. et al. (1990) FASEB J. 4, 2788–2797 69 Wang, H-J., Pisano, M.J. and Friedman, E. (1994) Neurobiol. Aging 15, 293–298 70 Masliah, E. et al. (1990) J. Neurosci. 10, 2113–2124 71 Nitsch, R.M. and Growdon, J.H. (1994) Biochem. Pharmacol. 47, 1275–1284 72 Govoni, S. et al. (1993) Neurology 43, 2581–2586 73 Bergamaschi, S. et al. (1995) Neurosci. Lett. 201, 1–4
Acknowledgements The authors wish to thank D. MochlyRosen for helpful comments, F.J. Van der Staay for providing the Wistar rats and W.C. Wetsel for the antisera to P.K.C. isoforms. This work was supported by the National Research Council (CNR) of Italy to F.B., the Ministry of University, Scientific and Technologic Research (MURST) to A.P., and by MURST and the Alzheimer Unit, Sacred Heart Hospital-FBF, Brescia, Italy to S.G.
Current views on eye development Guillermo Oliver and Peter Gruss Several genes involved in the regulation of eye development in different species have been identified. Structural and functional conservation have been found between some of these genes in organisms as diverse as Drosophila and mouse. One notable example is the relationship between the mouse Pax6 gene and eyeless of Drosophila. Ectopic expression of eyeless or mouse Pax6 in Drosophila results in the formation of additional eyes. Recently, another homeobox gene, Six3, was found to promote ectopic lens formation in fish embryos.The next step will be to unravel the associated regulatory pathways of these genes and assess the degree to which they display evolutionary conservation. This will be important in order to assimilate these findings with current anatomical and embryological models. It seems reasonable to believe that in the near future the characterization of the whole framework required for vertebrate eye development will be accomplished. Trends Neurosci. (1997) 20, 415–421
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N SCIENCE AND MYTHOLOGY the eye has always been an exciting topic. Many open questions still remain, particularly those concerning the evolutionary aspects of how different organisms have solved the problem of forming and detecting an image. The identification of various conserved genes among organisms as different as Drosophila and mice has challenged some Copyright © 1997, Elsevier Science Ltd. All rights reserved. 0166 - 2236/97/$17.00
of the previously accepted ideas regarding the origin of metazoan visual system. How much of the development pathway leading to eye formation has been conserved during evolution remains to be answered. Here, we focus on some of the most recent aspects of metazoan eye development, with a particular emphasis on the role of certain homeobox genes. PII: S0166-2236(97)01082-5
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Guillermo Oliver is at the Dept of Genetics, St Jude Children’s Research Hospital, 332 North Lauderdale, Memphis, TN 38105, USA. Peter Gruss is at the Dept of Molecular Cell Biology, MaxPlanck Institute of Biophysical Chemistry, Am Fassberg, 37077 Göttingen, Germany.
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Fig. 1. Structure of the compound eye of Drosophila. (A) Scanning electron micrograph of a wild-type fly eye. (B) Schematic view of an ommatidial unit. A longitudinal section is shown on the left and cross sections at three different levels are shown on the right. Abbreviations: A, photoreceptor cell axons; AC, anterior cone cell; B, bristle; C, liquid-filled pseudocone; CZ, cone cells; EQC, equatorial cone cell; L, lens; M, basal membrane; PC, posterior cone cell; PLC, polar cone cell; PP, primary pigment cell; Rh, rhabdomere; SP, secondary pigment cells; TP, tertiary pigment cells; 1–8, photoreceptor cells R1–R8. Reproduced, with permission, from E. Hafen and K. Basler.
Evolutionary considerations An important, as yet unresolved aspect of visual system evolution is whether the similarities found in different eye structures arise as a consequence of a common ancestor or due to evolutionary convergence. In most living and fossilized animals it is possible to identify some kind of light-sensitive organ1. These structures vary from simple eye-spots with a small number of receptors to well-developed eyes2. The analyses of the various types of eyes in different animals indicate that for the majority of eye’s functional parts there have been alternative but similar solutions. Based on structure, anatomic origins and phylogeny, Salvini and Mayr1 proposed that the lens-containing eye of a vertebrate and the compound eye of an insect are independent evolutionary developments, and that eyes might have evolved independently as many as 40 different times. The main sensory component of all eye structures are the photoreceptor cells. These cells are classified as either ciliary or rhabdomeric, but no phylogenetic correlation can be made based on this criterion. However,
Fig. 2. Early development of optic cup and lens in vertebrate embryos. (A) Optic grooves start to evaginate in the diencephalon towards the surface ectoderm. (B) Optic vesicles contact the overlying ectoderm, which then differentiates into lens placodes. (C) The optic vesicle and the lens placode invaginate. They will become the neural and pigmented retina and the lens vesicle respectively.
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the fact that light absorption and phototransduction seem to be mediated by a common mechanism has supported the idea that photoreceptor cells have a monophyletic origin3. All visual systems, including those from organisms as diverse as insects, cephalopods and vertebrates, share similar light receptor molecules called opsins3,4. Opsins are seven-transmembrane receptors linked to a vitamin A-derived chromophore that is responsible for absorbing photons. This process is mediated by a G protein-linked signaling cascade in all living organisms. The conservation of opsins among different organisms suggests that photoreceptor cells have originated from a common ancestor and that a simple ancestral photosensitive cell evolved into the different types of eyes we see today4. Importantly however, G protein-linked receptors similar to opsins are known to function not only in light detection but also in many other different physiological processes5. Strong support for a common monophyletic origin has come from the discovery of some key regulator genes involved in the control of similar developmental pathways in Drosophila and vertebrate eye formation.
Eye formation and lens induction The eye structures of vertebrates and arthropods are perfect examples of distinct optical solutions to the problem often regarded as receiving and integrating an image. The compound eye of Drosophila and the eye and lens of vertebrates are clearly different, and both have been studied extensively. However, recent findings suggest that despite these apparent differences, the formation of the arthropod and vertebrate eyes rely on a conserved developmental pathway. Visual detection in adult Drosophila relies on the two compound eyes and the three dorsal ocelli. The compound eye of the adult fly (Fig. 1) consists of a hexagonal array of ~800 individual units or ommatidia, each containing eight photoreceptor neurons (R1–R8), 12 accessory cells, lens-secreting cone cells and a mechanosensory bristle4. Initially, the eye develops during the larval stage from the eye imaginal disc: an epithelial monolayer specified in the late embryo. Photoreceptor cells differentiate in the posterior margin of the eye disc and appear progressively in a wave-like fashion over a period of two days. The anterior movement of this wave of neural differentiation is called the ‘morphogenetic furrow’6. During the larval period, differentiation starts with the formation of the morphogenetic furrow, which progresses from posterior to anterior across the disc epithelium. Anterior to the furrow are the dividing, undifferentiated progenitor cells; immediately behind the furrow, cells form differentiating clusters; and more posteriorly, these clusters acquire their final differentiated state. The vertebrate eye (Fig. 2) is a very complex structure that originates from primordial tissues derived from the wall of the diencephalon and the overlying surface ectoderm. A functional lens ultimately requires the accumulation of lens crystallins7. Lens induction is often cited as a classic example of embryonic induction. One of the first experiments to document an inductive effect was performed by Spemann in 1901 (Ref. 8). Using a hot needle, he destroyed the optic vesicle primordium of the frog
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Rana temporaria and found that the optic anlage was required for lens formation. The result was that lenses did not develop, leading Spemann to conclude that the optic vesicle is necessary for lens formation. Spemann’s conclusions were later confirmed by Lewis9, who removed an optic vesicle from the head of a frog embryo at the late neurula stage and transplanted it below the flank ectoderm in areas that normally do not form lenses. He observed lens formation in association with the transplanted retinal tissue, and argued that the optic rudiment was sufficient for lens induction10. Subsequent experiments started to raise doubts about the generality of the so-called ‘optic-cup’ model of lens induction. By repeating Spemann’s experiments, King11 found that lenses could form in the absence of contact with optic vesicles. Moreover, other researchers found that in many species lenses can form if retinal tissue is removed at the neural plate stage (reviewed in Ref. 12). Furthermore, transplant experiments in which host and donor tissue was labeled showed that the optic vesicle was not sufficient for lens induction12–14. Many studies have been carried out in Xenopus to try to explain the mechanisms of lens induction. By tissue grafting and labeling, a current working model has been proposed (reviewed in Ref. 10). Based on this model, it is thought that in Xenopus the anterior neural plate acts as an early inducer of lens ectoderm during the late gastrula stage, and that lens induction involves four phases: competence; bias; specification; and differentiation. Competence is the capacity of a specific tissue, in this case the head ectoderm, to respond to an inducing signal. For example, in Xenopus, the animal cap ectoderm exhibits different competences according to the developmental stage (mesodermal, neural or lens). Transplantation experiments performed by Henry and Grainger in Xenopus10,13 showed that embryonic head ectoderm is competent to respond to lens-inductive interactions very early in development, during the mid-gastrula stage. They also showed that this process depends on the developmental timer and not on specific tissue interactions10. Later, Servetnick and Grainger15 showed that only mid-to-late gastrula ectoderm is competent to respond to signals coming from the anterior neural plate. At this stage, a planar signal coming from the anterior neural plate region might provide the initial lens determination signal10. Another inductive signal might also be provided by the region of the endoderm that underlies the lens area and that ultimately gives rise to the foregut. The combination of these signals thus enables the competent region of the head ectoderm to become committed or biased to a lens-forming fate10. In Xenopus, this bias is thought to be established at the neural plate stage and this condition increases as induction takes place. At some point during development, enough lens induction has occurred to bias the ectoderm to form a lens without further induction. As development proceeds, the optic vesicle contacts the presumptive lens region at the end of neurulation. The optic vesicle does not seem to be critical for the early lens-inductive events; however, it might be required to position the lens within the head ectoderm and during the later stages of lens differentiation10, as lenses formed without an optic vesicle are very rudimentary.
Do different eyes share common genes and pathways? The majority of the models of eye evolution and eye formation are based on morphological and embryological data. Recently, however, a variety of molecular markers expressed in different parts of the metazoan visual system have been identified and characterized. These markers have allowed us to decipher some of the key players in the complex process of eye formation, and have raised many new and important questions. Drosophila, Caenorhabditis elegans, mouse and zebra fish have become the models of choice for those trying to correlate embryological and molecular aspects of development. One of the advantages of these systems is the availability of numerous natural or induced developmental mutations, including some in the visual system. Molecular and genetic analyses of eye development in Drosophila have largely concentrated on the cellular interactions that regulate cell fate determination in the late larval eye disc16. A lot of data are available on the development of the different photoreceptors (R1–R8). Receptor tyrosine kinases, epidermal growthfactor-like molecules and Ras signal transduction are all required during photoreceptor development (reviewed in Ref. 17). Genes that regulate early events in fly eye development, within and anterior to the morphogenetic furrow have also been identified. In a region anterior to this furrow cells have been found to express patterning genes16. Some Drosophila eye mutants, such as eyeless, eyes absent, sine oculis and eye gone, are characterized by the partial or complete absence of compound eyes. These mutations are thought to affect patterning genes that act in the initial events of the regulatory cascade3. In addition, some secreted molecules are known to regulate pattern formation in the eye imaginal disc. For example, Decapentaplegic is required for proliferation and initiation of pattern formation at the posterior edge of the eye disc; it has also been suggested that it mediates hedgehog function during progression of the morphogenetic furrow across the disc18. In mice, a variety of eye mutants is available19 and new mutations have been generated by means of homologous recombination technology. Saturation mutagenesis in zebra fish has produced a collection of mutants affecting almost every aspect of embryonic development. This has given rise to some interesting new eye phenotypes, and the affected gene products are now awaiting isolation20. At the same time, many genes expressed during the development of the visual system have been isolated in other organisms. As a result, genes of developmental importance in flies have been found to have counterparts in vertebrates, and in some cases the reverse is also true (Table 1). These previously unsuspected relationships between Drosophila and vertebrates have been uncovered in various developmental pathways. One of the best-known examples is the conserved role of the Hox genes in the patterning of the antero–posterior body axis21. Many different groups of genes are expressed during different stages of eye development, including various homeobox genes (reviewed in Refs 22,23). Some homeobox-containing genes seem to have a conserved and important function during the development of TINS Vol. 20, No. 9, 1997
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TABLE 1. Genes involved in eye development Gene Organismsa (mouse/Drosophila) Pax6/eyeless Pax2/NF Otx2/orthodenticle Six3/sine oculis Chox10/NF Prox1/prospero
Gain-of-functionb
Loss-of-functionc
Ascidian, chicken, frog, fruitfly, human, Drosophila (ectopic eyes) mouse, nematodes, nemertean, quail, rat, Xenopus (axial defects) zebra fish Mouse, human, zebra fish
eyeless/reduced eyes (Drosophila) Small eye (mouse, rat) Aniridia (humans) Optic nerve coloboma (KO mouse) Optic nerve coloboma (human mutation) Chicken, frog, fruitfly, human, mouse, sea Xenopus (extra cement Defective head development and no 55 urchin, zebra fish gland) ocelli (Drosophila)53 Microphthalmia/cyclopia (KO mouse) Fruitfly, mouse Medaka fish (ectopic lens) Eyeless/reduced eyes (Drosophila) Goldfish, mouse, nematode Ocular retardation (mouse) Chicken, frog, fruitfly, human, mouse, Axonal outgrowth and GMC nematode defects (Drosophila)
Eya/eyes absent
Fruitfly, human, mouse, nematode62
NF/dachshund
Fruitfly
Drosophila (ectopic eyes)
Eyeless/reduced eyes (Drosophila)61 Branchio–oto–renal syndrome (mouse) Reduced or not eyes (Drosophila)43
Abbreviations: GMC, ganglion mother cell; KO, knockout; NF, not found. a Organisms in which homolog genes were found. b The results observed in those cases in which gain-of-function experiments were performed in different organisms. c The results observed either in natural occurring mutations (italics) or in those generated by in vitro inactivation of the gene. In both cases, special emphasis is only given to the visual system phenotype. References are only given for those cases not previously mentioned in the text.
the metazoan visual system; they are transcription factors that, aside from being highly conserved in evolution, play fundamental roles during early embryonic pattern formation21. The best-known example is represented by the Pax6/eyeless genes. Pax6 is a member of the Pax family of genes and contains two DNA-binding domains, a paired-domain and a homeodomain. Initially, this gene was isolated from a mouse genomic library by using other previously identified murine Pax genes as molecular probes24. Pax6 was found to be expressed early in development in the anterior neural plate, optic vesicles, lens and nasal placodes25,26. Later studies showed that, in mammals, mutations in this gene were responsible for the mouse Small eye phenotype and the human genetic syndrome aniridia27,28. Homozygous Small eye mutant embryos have several defects, such as being born without eyes and nose, and die soon after birth. This mutation represents one of the first correlations between a well characterized spontaneous mutation and a specific molecular marker. Interestingly, in the homozygous mouse embryos, the optic vesicles start to develop but then degenerate27,28. Early in mammalian development, Pax6 is expressed in the surface ectoderm that will form the lens placode. In Xenopus, Pax6 has been shown to be expressed at the late gastrula stage in the anterior border of the developing neural plate and in the prospective retinal tissue29. In homozygous Small eye mutants, whereas the optic vesicles start to form, the lens placode never develops; consequently, no eyes are present. Tissue recombination experiments showed that ectoderm from homozygous rSey/rSey (rSey is the rat counterpart of the Sey mouse) will not differentiate into lens tissue once cultured with normal optic vesicles30. By contrast, lens differentiation occurs with wild-type ectoderm and mutant optic vesicle30. This suggests that in mutant animals, the ability of the ectoderm to respond to an inductive signal is affected and that Pax6 participates in the estab418
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lishment of the competence in the head ectoderm. Therefore, it is possible that in Small eye mutants the optic vesicle degenerates because an essential signal, probably emanating from the lens placode area, is missing. Furthermore, apparently functional Pax6 binding sites have been found within the Pax6 promoter31 and in the promoters and enhancers of several crystallin genes22,32. Homologous Pax6 genes were isolated from several other species including sea urchin33, nemertean34, ascidian35, chicken36, quail37 and zebra fish38. Surprisingly, the well-known eyeless mutation in Drosophila was due to a mutation in a Pax6-related gene. This gene was isolated from a Drosophila expression library using an oligonucleotide corresponding to a homeodomain binding site as a probe39. The chromosomal localization of the Pax6 homolog indicated that it corresponded to the Drosophila eyeless mutation39. The eyeless gene is expressed in eye progenitor cells and during differentiation; it is strongly expressed anterior to and within the morphogenetic furrow39; and is also expressed during Drosophila nervous system development. Homozygous eyeless flies show a reduction or lack of eyes. These findings indicate that despite the clear anatomical differences between the compound Drosophila eye and the vertebrate eye, some conservation at the gene level has been maintained. Further support for this idea comes from the report of induction of ectopic eyes in Drosophila by targeted expression of either the fly eyeless gene product or the mouse Pax6 gene in the wings, legs and antennae imaginal discs40. However, misexpression of Pax6 during early Xenopus development resulted in axial defects, but no specific eye phenotype29. These results, together with the isolation of Pax6related genes in such evolutionary distant species as nemertines, ascidians, sea urchin and nematodes3, strongly suggest that Pax6/eyeless is one of the major
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players in visual system development in metazoans3,40. Interestingly, the nematode C. elegans, in which a Pax6 homolog gene has been identified41,42 has no eyes. In this case, Pax6 is involved in the patterning of the head region and it is also required for the development of a tail sense organ41,42. Targeted mis-expression has been performed in Drosophila with the product of the dachshund gene43. This gene encodes a nuclear protein required for normal eye formation in the fly. Similar to the result obtained with eyeless, mis-expression of dachshund induced ectopic eye formation in Drosophila43. Furthermore, the authors showed that dachshund and eyeless induce the expression of each other, and that dachshund is required for ectopic retinal development driven by eyeless mis-expression. No vertebrate counterpart for dachshund has yet been identified. Pax2, another member of the Pax family of transcription factors is also expressed during murine eye development44,45. Its expression is confined to those cells within the optic vesicle that contribute to the optic stalk. Pax2 knockout mice have coloboma, a failure of the optic fissure to close, indicating that this gene is required for closure of the fissure45. Human46 and zebra fish47 Pax2 homologs have also been identified. A Pax2 natural mutation in humans was found to be associated with abnormal choroid fissure formation46. Experiments with zebra fish homologs of these two Pax genes have shown that two members of the hedgehog family of signaling molecules, normally expressed in the midline, can promote Pax2 expression and inhibit Pax6 within the optic vesicle48,49. These data suggest that the hedgehog proteins, directly or indirectly, regulate Pax2 and Pax6 expression. Recent studies also indicate that a single morphogenetic field gives rise to the two separate retina primordia under the influence of the prechordal plate50. Sonic hedgehog, a gene expressed in the midline, seems a strong candidate for this retina patterning, probably by suppressing Pax6 expression in the medial part of the neural plate50. In Drosophila, the hedgehog gene is expressed in differentiating cluster cells but also influences events anterior to the morphogenetic furrow. Flies mutant for this gene show reduced eyes and cell death. Two vertebrate homologs of the Drosophila homeobox-containing gene orthodenticle have been isolated (Otx1 and Otx2) and both are expressed in the head and brain of the developing mouse embryo51. However, both genes are also expressed in the developing nasal epithelium and in different parts of the developing visual system51. Disruption of the Otx2 gene in mice causes microphthalmia, a phenotype associated with the eyes either protruding over the face surface or buried in the orbit. The retina is hyperplastic, and in some cases only one eye is present, usually without lens and cornea52. This suggests that Otx2 participates in some stages of eye development possibly by specifying anterior head formation. In Drosophila, orthodenticle is expressed in the most anterior part of the head, and it is also required for the formation of the ocelli in the adult fly53. Otx2-related genes were also identified in sea urchin, zebra fish, humans (see Ref. 54), Xenopus55 and chicken56. Chx10 is another murine homeobox gene expressed in the optic vesicles and neuroretina, particularly in
the inner nuclear layer57. Homologous genes were isolated in goldfish and C. elegans58. A naturally occurring mouse mutation in this gene was shown to be responsible for the ocular retardation phenotype (reduced cell proliferation rate in the neural retina)57. These data support the idea that Chx10 is essential for retinal cell proliferation and for bipolar cell and rod photoreceptor differentiation59. Three murine homologs of the Drosophila eyes absent gene have been isolated60; in flies, the eyes absent gene is expressed anterior to the furrow and encodes a novel nuclear protein required for eye development61; the murine homologs Eya1 and Eya2 are expressed during the development of the cranial placodes, and all three are expressed in the developing eye60. Mutations in the human Eya1 gene have been found to be responsible for the branchio–oto–renal syndrome62. Another interesting parallel between Drosophila and vertebrate eyes is represented by the prospero/Prox1 genes. The product of the fly prospero gene is expressed in the lens-secreting cone cells63, whereas its putative murine homolog, Prox1 is expressed in the lens fibers64. This suggests that lens development in different species shares some common conserved molecular mechanisms. Prospero-related genes have also been identified in C. elegans65, chicken66, human67 and Xenopus (G. Oliver, unpublished). The product of the Drosophila sine oculis gene (a homeobox-containing gene) is expressed anterior to the furrow and it is essential for the development of the entire visual system by controlling the initial events of pattern formation in the eye disc16. The sine oculis gene is expressed before patterning and its product is required for formation of the optic lobes. In some sine oculis mutants, ommatidia fail to develop, suggesting that this gene is essential for the initiation of pattern formation in the disc. These mutant flies also exhibit apoptosis anterior to the morphogenetic furrow and their optic lobe primordia do not invaginate16. Recently, a mammalian gene family with high sequence similarity to the sine oculis fly gene was identified68. This family contains at least five members, one of which, Six3, seems to be the murine functional homolog of sine oculis69. Interestingly, in mouse, the expression of some of the Six genes co-localizes in many tissues with that of Eya1 and Eya2 (Refs 60,68,69). Six3 is expressed in the anterior neural plate and prospective eye field (Fig. 3), optic stalk, optic vesicles, lens and nasal placode69. This expression pattern, together with the fact that the anterior neural plate is involved in Xenopus lens induction10, suggests that Six3 has a role in vertebrate eye formation. In an attempt to understand this role, we ectopically expressed mouse Six3 in fish embryos70. We found that Six3 was sufficient to promote ectopic lens formation in the area of the otic vesicle (Fig. 4) and that retinal tissue is not a prerequisite for ectopic lens differentiation. The finding that ectopic expression of a single mouse gene (Six3) promotes lens formation in a fish embryo demonstrates that this gene is a key player in this process70. The results also suggest that, like Pax6 in vertebrates and insects, Six3/sine oculis is also evolutionarily conserved in metazoan eye development. The lens placode is one of the sensory placodes that is represented by thickening of the head ectoderm surrounding the neural plate. In addition to the lens TINS Vol. 20, No. 9, 1997
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Fig. 3. (Left) Six3 early expression in the prospective eye field. Whole mount in situ hybridization of an 8.5-day-old mouse embryo (ventral view). Purple staining indicates Six3 expression in the anterior neural plate region. Eyes will later develop in that region. Fig. 4. (Right) Ectopic Six3 expression promotes lens formation in the otic vesicle territory. Three-day-old medaka embryo showing an ectopic lens (arrow) replacing the left otic vesicle (OV). The lens (L), lens epithelium (arrowhead) and neuroretina (NR) of the wild-type eyes are shown. Whole mount in situ hybridization analysis was performed on this embryo leading to an unspecific brown staining of the lenses.
placode, the other sensory placodes are: the nasal placode, which gives rise to the nasal epithelium; and the otic placode, which forms the inner ear. Jacobson proposed that initially a common equipotential placodal state is activated in a large region of head
ectoderm, and later, via different interactions with the developing brain, the specific sensory placodes form71. The induction of ectopic lenses in the otic placode territory demonstrates that the region of the otic placode is competent to form lenses in response to Six3, and supports previous tissue transplantation data72. Based on the proposed model for lens induction discussed above10, it is possible that ectopic Six3 expression changes the bias of the otic placode towards the lens pathway by inducing the secretion of some unknown factor70 (Fig. 5). In this case, an ectopic lens could form in the area in which the inner ear would normally develop. It is possible that during early development, the whole head ectoderm is competent to respond to a variety of different signals. The establishment of an unreversible commitment (bias) towards optic, otic or nasal placodes would thus depend on which signal or signals were available. In this scenario, Six3 could play a crucial role in the pathway of lens placode determination (Fig. 5). By analogy, one might predict that genes will be identified that can induce otic or nasal structures when expressed ectopically in uncommitted head ectoderm. To confirm the function of the Six3 gene in eye development we are currently working on the generation of Six3 null-mutant mice. Finally, it is still not yet clear whether the different metazoan eyes have evolved from a common ancestor or whether they have evolved independently on many different occasions using generic signaling pathways – in this case, similar molecules were recruited for this purpose. It is possible that some of the genes discussed previously (Pax6, Otx2, Six3 and Eya) were initially involved in the patterning of the anterior part of the embryo. The identification of a Pax6 homolog in an organism such as C. elegans, devoid of eyes, and its suggested role in the patterning of the anterior head structures also supports this proposal. Further on in evolution, the aforementioned genes become involved in the induction and differentiation of cranial placodes; in this respect, not only are they similarly expressed in anterior forebrain and eyes, but also in the nasal placodes and nasal epithelium, and some members of this group are also expressed in the otic placode and pituitary. Later, these and other still-unidentified genes might have been recruited to regulate different conserved structural molecules involved in light or other stimulus detection. From there, the different types of metazoan eyes might have evolved. It will be extremely interesting to find out whether the regulatory cascades in which these genes function have also been conserved during the evolution of eye development. Selected references
Fig. 5. Proposed model of ectopic lens induction by ectopic Six3 expression in fish embryos. According to the current model for lens induction in Xenopus10, in the wild-type situation, early in development the head ectoderm is still uncommitted. Later, and via different molecules, a competent phase for each of the sensory placodes is acquired. Finally, each of the placodes becomes unreversibly biased: nose, internal ears or lens will be specified and they will differentiate. In the case of the transgene, the ectopic overexpression of Six3 has probably affected the transition from competence to bias. As a result, what could eventually be considered a homeotic transformation occurred, and a lens was induced instead of the otic placode. Abbreviations: Fb, forebrain; Hb, hindbrain; Mb, midbrain.
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1 Salvini-Plawen, L.V. and Mayr, E. (1977) Evol. Biol. 10, 207–263 2 Land, M.F. and Fernald, R.D. (1992) Annu. Rev. Neurosci. 15, 1–29 3 Halder, G., Callaerts, P. and Gehring, W.J. (1995) Curr. Opin. Genet. Dev. 5, 602–609 4 Zuker, C.S. (1994) Science 265, 742–743 5 Nilsson, D.E. (1996) Curr. Biol. 6, 39–42 6 Wolff, T. and Ready, D.F. (1993) in The Development of Drosophila melanogaster (Bate, M. and Martinez-Arias, A., eds), pp. 1277–1326, Cold Spring Harbor Laboratory Press 7 Wistow, G. (1993) Trends Biochem. Sci. 18, 301–306 8 Spemann, H. (1901) Verb. Anat. Ges. 15, 61–79 9 Lewis, W. (1904) Am. J. Anat. 3, 505–536 10 Grainger, R.M. (1992) Trends Genet. 8, 349–355 11 King, H.D. (1905) Arch. Entw. Mech. 19, 85–107 12 Saha, M.S., Spann, C.L. and Grainger, R.M. (1989) Cell Diff. Dev. 28, 153–172 13 Henry, J. and Grainger, R. (1987) Dev. Biol. 124, 200–214
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14 Grainger, R., Henry, J. and Henderson, R. (1988) Development 102, 517–526 15 Servetnick, M. and Grainger, R. (1991) Development 112, 177–188 16 Cheyette, B.N.R. et al. (1994) Neuron 12, 977–996 17 Zipursky, L. and Rubin, G. (1994) Annu. Rev. Neurosci. 17, 373–397 18 Pignoni, F. and Zipursky, L. (1997) Development 124, 271–278 19 Graw, J. (1996) Dev. Genet. 18, 181–197 20 Malicki, J. et al. (1996) Development 123, 263–273 21 De Robertis, E.M., Oliver, G. and Wright, C.V.E. (1990) Sci. Am. 262, 46–52 22 Cvekl, A. and Piatigorsky, J. (1996) BioEssays 18, 621–630 23 Beebe, D.C. (1994) Invest. Ophthalmol. Vis. Sci. 35, 2897–2900 24 Walther, C. et al. (1991) Genomics 11, 424–434 25 Walther, C. and Gruss, P. (1991) Development 113, 1453–1449 26 Grindley, J.C. et al. (1995) Development 121, 1433–1442 27 Hill, R.E. et al. (1991) Nature 354, 522–525 28 Ton, C.C.T. et al. (1991) Cell 67, 1059–1074 29 Hirsch, N. and Harris, W.A. (1997) J. Neurobiol 32, 45–61 30 Fujiwara, M. et al. (1994) Differentiation 57, 31–38 31 Plaza, S. et al. (1995) Mol. Cell. Biol. 15, 892–903 32 Richardson, J., Cvekl, A. and Wistow, G. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 4676–4680 33 Czerny, T. and Busslinger, M. (1995) Mol. Cell. Biol. 15, 2858–2871 34 Loosli, F., Kmita-Cunisse, M. and Gehring, W.J. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 2657–2663 35 Glardon, S. et al. (1997) Development 124, 817–825 36 Goulding, M.D., Lumsden, A. and Gruss, P. (1993) Development 117, 1001–1016 37 Dozier, C. et al. (1993) Cell Growth Differ. 4, 281–289 38 Krauss, S. et al. (1991) EMBO J. 10, 3609–3619 39 Quiring, R. et al. (1994) Science 265, 785–789 40 Halder, G., Callaerts, P. and Gehring, W.J. (1995) Science 267, 1788–1792 41 Chisholm, A.D. and Horvitz, H.R. (1995) Nature 377, 52–55
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Acknowledgements We thank S. Vallance, T. Curran and L. Zipursky for critical reading of the manuscript, M. Kessel for helping in the preparation of Fig. 5, and an anonymous reviewer for some helpful suggestions.
Development of the insect stomatogastric nervous system Volker Hartenstein The stomatogastric nervous system (SNS) forms a network of peripheral ganglia associated with the insect gut.The SNS originates from a neuroepithelial placode which dissolves into a population of migrating neural precursors. The formation of the SNS presents many parallels to the development of the vertebrate peripheral nervous system. Recent studies have started to provide answers for pertinent questions in SNS development,in particular,how the SNS placode is specified, how SNS precursors are released in a reproducible pattern from this placode and how different cell types in the SNS are determined. Trends Neurosci. (1997) 20, 421–427
C
OMPARABLE IN FUNCTION to the autonomic nervous system of vertebrate organisms, there exists in invertebrates a system of peripheral ganglia and nerve fibres that innervate the visceral organs and control food digestion, respiration and excretion. In higher invertebrates, including annelids and arthropods, the anterior portion of the visceral nervous system surrounding the foregut consists of a chain of ganglia which are connected in a characteristic manner with the brain and endocrine system. These nerve centres innervate muscles of the mouth cavity, foregut and midgut, and regulate food uptake and food transport; they are referred to as the stomatogastric nervous system (SNS). Due to its small cell number, simple behavioral output and easy accessibility, the SNS of crustaceans has
Copyright © 1997, Elsevier Science Ltd. All rights reserved. 0166 - 2236/97/$17.00
been used for many years as a model system to study the functioning of a neural network1. More recently, the development of the stomatogastric nervous system in insects has attracted the attention of several investigators2–9, in part due to the fact that in some aspects it resembles neural development in vertebrates. The vertebrate autonomic nervous system is derived from the neural crest. During neurulation, the neurectoderm (neural plate) invaginates and forms the neural fold. Most of the neural fold closes dorsally to form the neural tube (giving rise to the CNS); cells at the lateral fringes of the fold move out of the epithelium and become the neural crest10,11. Subsequently, these cells migrate along different routes and form the various ganglia of the sensory and autonomic nervous PII: S0166-2236(97)01066-7
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Volker Hartenstein is at the Dept of Molecular, Cell and Developmental Biology, University of California, Los Angeles, Los Angeles, CA 90095-1606, USA.
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