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					Current Biology Vol 15 No 19 R796

equally matched by their spectacular diversity in morphology and body colouration [10]. As well as expanding into unoccupied ecological niches, sexual selection of body colour variation among males, by female cichlids, is thought to contribute to the incredibly fast rates of speciation [10]. Sexual selection may be an even more important factor in driving rapid speciation in the clear-water environment of Lake Malawi. It would be very interesting to compare the spectral composition of body colour patterns with the maximal spectral sensitivities of the expressed cone opsins in the Lake Malawi cichlids. Selection pressure to match, or tune, the visual sensitivities of the opsins to the local light environment may also contribute to speciation. In the area of visual ecology, there are many well-established correlations between spectral composition of the light environment and maximal spectral sensitivities of visual pigments [11]. For example, deep-sea fish live in a short wavelength-rich light environment, which is matched by a short wavelength shifted maximal sensitivity of their Rh1 opsin and rod photoreceptor [11]. Careful analysis of the opsin genes of the Lake Malawi (clear water, short wavelength-rich) and Lake Victoria (turbid water, long wavelength-rich) cichlids has revealed positive selection in opsin genes in species living in the two different environments [12]. This is important, as it takes the numerous observations of matched light environment and visual sensitivities beyond a series of interesting correlations and demonstrates positive selection and adaptive, photic environmentdriven evolution in the East African cichlids. From genome sequencing data, the rate of gene duplication has been estimated at 0.01 duplications per gene per million years [8]. This is much higher than the observed number of duplicated genes because gene gain is generally balanced by a short half-life of new genes and a high rate of gene loss [8]. The identification of seven cone opsin

genes [9] and therefore the potential for heptachromatic vision in the Lake Malawi cichlids opens up a Pandora’s box of untapped evolutionary potential that is constantly creating new genes. The present evolutionary circumstances of the cichlid fish populations in the East African Lakes is providing a window into how rapidly molecular evolutionary mechanisms can kick into top gear, switch the balance towards keeping those newly duplicated genes and exploring new areas of genetic space. This type of ‘evolution in waiting’ may ultimately govern the expansion and diversification of whole populations to exploit newly available ecological niches through rapid speciation.
1. Darwin, C.R. (1859). The Origin of Species by Means of Natural Selection. (New York: Gramercy Books.) 2. Ohno, S. (1970). Evolution by Gene Duplication. (New York: Springer-Verlag.) 3. Klare, J.P., Gordeliy, V.I., Labahn, J., Buldt, G., Steinhoff, H.J., and Engelhard, M. (2004). The archaeal sensory rhodopsin II/transducer complex: a model for transmembrane signal transfer. FEBS Lett. 564, 219–224. 4. Dawkins, R. (2004). The Ancestor’s Tale: A Pilgrimage to the Dawn of Life. (London: Weidenfeld & Nicolson.)

5. Ota, S. and Saitou, N. (1999). Phylogenetic relationship of muscle tissues deduced from superimposition of gene trees. Mol. Biol. Evol. 16, 856–867. 6. Yokoyama, S. (2000). Molecular evolution of vertebrate visual pigments. Prog. Retin. Eye Res. 19, 385–419. 7. Collin, S.P., Knight, M.A., Davies, W.L., Potter, I.C., Hunt, D.M., and Trezise, A.E. (2003). Ancient colour vision: multiple opsin genes in the ancestral vertebrates. Curr. Biol. 13, R864–R865. 8. Lynch, M., and Conery, J.S. (2003). The origins of genome complexity. Science 302, 1401–1404. 9. Parry, J.W.L., Carleton, K.L., Spady, T., Carboo, A., Hunt, D.M., and Bowmaker, J. (2005). Mix and match colour vision: tuning spectral sensitivity by differential opsin gene expression in Lake Malawi cichlids. Curr. Biol. 15, this issue. 10. Salzburger, W., Mack, T., Verheyen, E., and Meyer, A. (2005). Out of Tanganyika: genesis, explosive speciation, keyinnovations and phylogeography of the haplochromine cichlid fishes. BMC Evol. Biol. 5, 17. 11. Bowmaker, J.K. (1995). The visual pigments of fish. Prog. Ret. Eye Res. 15, 1–31. 12. Spady, T.C., Seehausen, O., Loew, E.R., Jordan, R.C., Kocher, T.D., and Carleton, K.L. (2005). Adaptive molecular evolution in the opsin genes of rapidly speciating cichlid species. Mol. Biol. Evol. 22, 1412–1422.

School of Biomedical Sciences, University of Queensland, St Lucia, Brisbane, Qld. 4072, Australia. E-mail:
DOI: 10.1016/j.cub.2005.09.025

Axon Pruning: C. elegans Makes the Cut
Axon pruning has recently been described in the simple nervous system of the nematode Caenorhabditis elegans. Generating excess processes and pruning may be a phylogenetically conserved feature reflecting a flexibility to modify neural circuits. William G. Wadsworth Axon pruning is a means to modify the patterning of axons in a nervous system. For example, during embryonic development a transient nervous system arises that may contain excessive and unnecessary projections. Axon pruning is one means to eliminate inappropriately made connections and to help transform the nervous system into a mature state containing specific patterns of neural circuits [1]. This pruning of axons has been observed in different organisms when the nervous system becomes remodeled. During the development of layer V of the mammalian cortex, for example, different classes of neurons initially send axon branches to both the spinal cord and the superior colliculus. Later, the motor cortical neurons selectively prune their branches to the superior colliculus, while visual cortical neurons selectively prune their branches to the spinal cord [2] (Figure 1A). Axon pruning is also observed in holometabolous insects such as Drosophila and Manduca sexta. In such insects the nervous system is extensively reorganized during

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metamorphosis to generate an adult nervous system that is capable of mediating the distinct behaviors of the adult. In the embryonic and early larval stages, the γ neurons of the Drosophila mushroom bodies send axons into the larval medial and dorsal mushroom body axon lobes. During metamorphosis these neurons prune their axon branches and then regrow just the medial axon branch [3] (Figure 1B). Likewise, synaptic partners of the γ neurons also undergo stereotypic pruning of their axon branches during metamorphosis [4]. New findings published recently in Current Biology by Kage et al. [5] suggest that axon pruning also occurs in the nematode Caenorhabditis elegans. The AIM neurons are a set of two interneurons with cell bodies on the left and right sides of the ventral ganglion near the pharynx. In the mature nervous system their processes run anteriorly from the cell bodies. In Kage et al.’s study [5] it was observed that an additional extension connects the two AIM neurons in nearly 70% of L1 stage larvae and that the number of animals with this connection declines to approximately 10% in the L2, L3, L4, and adult stages (Figure 1C). It may seem surprising that excess processes might be produced and then pruned in C. elegans, because the hermaphrodite nervous system of C. elegans is relatively simple with each of the 302 neurons having a highly reproducible morphology and characteristic synaptic connections [6]. Thus, there are no large pools of neurons that project to the same targets, as is the case in vertebrates, and, consequently, no need to eliminate the excessive projections. Furthermore, by the end of embryogenesis the pattern of the C. elegans adult nervous system is laid down and the type of repatterning seen in holometabolous insects is not required. The common feature of pruning in C. elegans and the other organisms mentioned above is that it occurs as part of a

A Select



C Trim

Current Biology

Figure 1. Pruning cuts. (A) Selective elimination of branches can create specific neuronal groups. For example, layer V cortical projections are segregated to different subcortical targets in mammals. (B) Connections can be reorganized. For example, the γ neurons in the Drosophila mushroom bodies regrow and can develop different patterns following metamorphosis. (C) Excess projections can be trimmed. For example, an extra projection is removed from the AIM neurons in C. elegans.

transitional process. While most cells of the C. elegans nervous system arise during embryogenesis, additional cells are added during the L1 stage [7]. New motor neurons join the ventral nerve cord and there is remodeling of the synaptic output of some of the embryonic-derived motor neurons [8]. Also, during this period molecules known to influence nervous system development change their expression patterns or are first expressed. For example, a new expression pattern of the axon guidance molecule UNC-6/netrin emerges as the newly produced motor neurons express the protein [9]. First detected at this stage are the ZIG proteins, which help maintain the organization of the mature nervous system [10]. The timing of the synaptic remodeling and of zig gene expression are known to be controlled by lin-14, a hetrochronic gene that also regulates the timing of stagespecific cell division patterns [11,12]. Kage et al. [5] show that AIM pruning is regulated by phylogenetically conserved factors. Mutations in the mbr-1 and unc-86 genes impair the axon pruning, causing more of the extensions to be present in adults. The mbr-1 gene encodes a new member of a family of transcription factor proteins that includes orthologues in other invertebrates and in vertebrates [5]. UNC-86 is a POU-domain transcription factor that is important for generating asymmetry in the neuroblast cell lineages and for allowing diverse

neural types to develop [13]. Mammalian orthologs of UNC-86 have important roles in the development and function of neurons [14]. In the AIM neurons, UNC-86 may regulate mbr-1 expression [5] as well as the synthesis of the neurotransmitter serotonin, which is first detected at the L1 stage [15]. Serotonin production is associated with the terminal differentiation of the neurons [15], suggesting that regulation of the pruning is coupled to the process that determines the final identity of the neurons. The significance of the AIM pruning is unknown and whether the excess branch impacts behavior has not been determined. It is interesting that even within the wild-type population, some individuals retain the extra AIM branch. This raises a question; to what degree does variation exist in the axon patterns of C. elegans? This issue has not been systematically studied, but other neurons are known to have variable axon patterns; for example, a branch of the PDE axon is short or missing in about 5% of wild-type animals [16]. There is evidence that axon morphology during larval development is sensitive to activity [17] and an intriguing possibility is that experience and neuronal activity might influence pruning and patterning of the axons, allowing some variation in neural circuit patterns within a population. The discovery of excess processes and axon pruning

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during the development of a relatively simple nervous system is provocative. The ability to tolerate extra axon processes and to modify connections postembryonically by pruning and other means could be a phylogenetically conserved characteristic of nervous system development. This trait might allow greater flexibility to adopt behaviors suitable to changing environmental conditions.
1. Kantor, D.B., and Kolodkin, A.L. (2003). Curbing the excesses of youth: molecular insights into axonal pruning. Neuron 38, 849–852. 2. O’Leary, D.D., and Koester, S.E. (1993). Development of projection neuron types, axon pathways, and patterned connections of the mammalian cortex. Neuron 10, 991–1006. 3. Lee, T., Lee, A., and Luo, L. (1999). Development of the Drosophila mushroom bodies: sequential generation of three distinct types of neurons from a neuroblast. Development 126, 4065–4076. 4. Marin, E.C., Watts, R.J., Tanaka, N.K., Ito, K., and Luo, L. (2005). Developmentally programmed remodeling of the Drosophila olfactory circuit. Development 132, 725–737. 5. Kage, E., Hayashi, Y., Takeuchi, A.,








Hirotsu, T., Kunitomo, T.I., Arai, H., Iino, Y., and Kubo, T. (2005). MBR-1, a novel helix-turn-helix transcription factor, is required for pruning excessive neurites in Caenorhabditis elegans. Curr Biol. 15, 1554–1559. White, J., Southgate, E., Thompson, J., and Brenner, S. (1986). The structure of the nervous system of the nematode Caenorhabditis elegans. Philos. Trans. R. Soc. Lond. B Biol. Sci. 314, 1–340. Sulston, J.E., and Horvitz, H.R. (1977). Post-embryonic cell lineages of the nematode, Caenorhabditis elegans. Dev. Biol. 56, 110–156. White, J.G., Alberson, D.G., and Annes, M.A.R. (1978). Connectivity changes in a class of motorneurons during the development of a nematode. Nature 271, 764–766. Wadsworth, W.G., Bhatt, H., and Hedgecock, E.M. (1996). Neuroglia and pioneer neurons express UNC-6 to provide global and local netrin cues for guiding migrations in C. elegans. Neuron 16, 35–46. Aurelio, O., Hall, D.H., and Hobert, O. (2002). Immunoglobulin-domain proteins required for maintenance of ventral nerve cord organization. Science 295, 686–690. Aurelio, O., Boulin, T., and Hobert, O. (2003). Identification of spatial and temporal cues that regulate postembryonic expression of axon maintenance factors in the C. elegans ventral nerve cord. Development 130, 599–610. Hallam, S.J., and Jin, Y. (1998). lin-14 regulates the timing of synaptic remodelling in Caenorhabditis elegans. Nature 395, 78–82.






Baumeister, R., Liu, Y., and Ruvkun, G. (1996). Lineage-specific regulators couple cell lineage asymmetry to the transcription of the Caenorhabditis elegans POU gene unc-86 during neurogenesis. Genes Dev. 10, 1395–1410. McEvilly, R.J., and Rosenfeld, M.G. (1999). The role of POU domain proteins in the regulation of mammalian pituitary and nervous system development. Prog. Nucleic Acid Res. Mol. Biol. 63, 223–255. Sze, J.Y., Zhang, S., Li, J., and Ruvkun, G. (2002). The C. elegans POU-domain transcription factor UNC-86 regulates the tph-1 tryptophan hydroxylase gene and neurite outgrowth in specific serotonergic neurons. Development 129, 3901–3911. Hedgecock, E.M., Culotti, J.G., and Hall, D.H. (1990). The unc-5, unc-6, and unc40 genes guide circumferential migrations of pioneer axons and mesodermal cells on the epidermis in C. elegans. Neuron 4, 61–85. Peckol, E.L., Zallen, J.A., Yarrow, J.C., and Bargmann, C.I. (1999). Sensory activity affects sensory axon development in C. elegans. Development 126, 1891–1902.

UMDNJ-Robert Wood Johnson Medical School, Department of Pathology, 675 Hoes Lane West, Piscataway, New Jersey 08854-5635, USA. E-mail:
DOI: 10.1016/j.cub.2005.09.024

Intraflagellar Transport: Keeping the Motors Coordinated
Intraflagellar transport is a conserved delivery system that services eukaryotic cilia and flagella. Recent work in the nematode Caenorhabditis elegans has identified proteins required for the functional coordination of intraflagellar transport motors and their cargoes. Douglas G. Cole Eukaryotic cilia and flagella are remarkable machines that serve a variety of sensory and motile functions. Interest in these organelles has expanded recently as their assembly and function have become associated with a number of human diseases, including polycystic kidney disease and Bardet-Biedl syndrome [1,2]. Essential for the construction of cilia and flagella is the process known as intraflagellar transport or IFT. First identified in the green alga Chlamydomonas reinhardtii [3], IFT consists of long trains of proteinaceous particles that are moved out to the distal tip by kinesin-2 (anterograde IFT) and moved back toward the cell body by cytoplasmic dynein 1b/2 (retrograde IFT) [4,5]. In turn, anterograde IFT particles supply precursors that are used to assemble the axonemal cytoskeleton, while retrograde IFT particles are responsible for removal of turnover products [6]. IFT is also required for the directed movement of other ciliary proteins including the PKDassociated qilin [7] and specific membrane channels [8]. In the nematode, Caenorhabditis elegans, two related kinesins combine forces to drive anterograde IFT along

sensory cilia [9]. Identified in a screen of mutants defective in osmotic avoidance, the homodimeric OSM-3 is exclusively expressed in a subset of ciliated neurons responsible for chemosensation [10]. Expressed in all ciliated neurons is the heterotrimeric kinesin-2, which contains two unique motor subunits, KLP-11 and KLP-20, and a third, cargo-adaptor subunit known as the kinesin-associated polypeptide or KAP [9]. The dendritic sensory cilia of the nematode consist of three sections, with a 1 µm proximal segment that is the functional equivalent of a transition zone. The middle segment contains 4 µm doublet microtubules that lead to a distal segment of 2.5 µm singlet microtubules. Kinesin-2 and OSM-3 coordinate to jointly move IFT particles at 0.7 µm sec–1 in the middle segments, while OSM-3 alone moves the particles at a faster rate of 1.3 µm sec–1 along the singlet microtubules of the distal segment (Figure 1A) [11].

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