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Ultrastructural and dye filling studies have suggested that GABAergic interneurons make gap junctions onto mitral cells in the mammalian bulb [14,15]. However, this remains a controversial topic that requires further work. New Questions The study by Zhu et al. [1] raises many questions. First, exactly which interneurons are involved? The dlx4/6 neurons are a heterogeneous population [1], like the interneurons of the Drosophila antennal lobe, and some of these neurons may play entirely distinct roles. Identifying a genetic marker for each relevant interneuron type would permit more targeted recordings, better mapping of connectivity, and more precise manipulations. It is also notable that there are likely many other GABAergic interneurons in the fish olfactory bulb in addition to the dlx4/6 neurons. In the mammalian bulb there are several populations of periglomerular cells in the superficial glomerular layer, as well as deep-layer granule cells, which outnumber all other cell types in the bulb by at least an order of magnitude [16]. What specific roles do each of these different inhibitory circuits play, and how do they interact? Second, do specific odor stimuli elicit specific spatial patterns of interactions among olfactory glomeruli? Or alternatively, is this a global interaction that simply scales in strength with the total level of afferent input to the circuit? None of the studies discussed here provides a clear answer to this question. This issue is critical to understanding how interneurons affect olfactory processing. Third, why does this circuit have such diverse effects on different target neurons? When the dlx4/6 neurons were manipulated, there were large variations across cells in the effects this had on neural activity. For a given odor stimulus, some cells were inhibited, but others were disinhibited [1]. Again, this finding has a parallel in the Drosophila antennal lobe, where identified olfactory glomeruli have diverse levels of sensitivity to lateral inhibition and lateral excitation [8,13,17]. What are the mechanisms and significance of this diversity? Finally, how deep are the functional and structural similarities between neural circuits in different organisms? We should remember that the success
of molecular biology in the 20th century hinged on our ability to spot homologies between amino acid sequences in different protein domains, and to grasp the systematic relationship between the primary structure of a domain and its function. Many such molecular modules are now known to reoccur throughout the animal kingdom. Being able to spot similar kinds of structure–function relationships in neural circuits would accelerate discovery in neuroscience. For this reason, comparative biology remains essential to the search for general principles. References 1. Zhu, P., Frank, T., and Friedrich, R.W. (2013). Equalization of odor representations by a network of electrically coupled inhibitory interneurons. Nat Neurosci. 16, 1678–1686. 2. Fairhall, A. (2014). Adaptation and natural stimulus statistics. In The Cognitive Neurosciences 5th Edition, M.S. Gazzaniga, ed. (Cambridge, MA: MIT Press). 3. Laughlin, S.B. (2001). Energy as a constraint on the coding and processing of sensory information. Curr. Opin. Neurobiol. 11, 475–480. 4. Shapley, R., and Enroth-Cugell, C. (1984). Visual adaptation and retinal gain controls. Prog. Retin. Res. 3, 263–346. 5. Carandini, M., and Heeger, D.J. (2012). Normalization as a canonical neural computation. Nat. Rev. Neurosci. 13, 51–62. 6. Zoccolan, D., Cox, D.D., and DiCarlo, J.J. (2005). Multiple object response normalization in monkey inferotemporal cortex. J. Neurosci. 25, 8150–8164. 7. Friedrich, R.W., and Korsching, S.I. (1997). Combinatorial and chemotopic odorant coding in the zebrafish olfactory bulb visualized by optical imaging. Neuron 18, 737–752. 8. Olsen, S.R., Bhandawat, V., and Wilson, R.I. (2010). Divisive normalization in olfactory population codes. Neuron 66, 287–299. 9. Asahina, K., Louis, M., Piccinotti, S., and Vosshall, L.B. (2009). A circuit supporting concentration-invariant odor perception in Drosophila. J. Biol. 8, 9.
10. Olsen, S.R., and Wilson, R.I. (2008). Lateral presynaptic inhibition mediates gain control in an olfactory circuit. Nature 452, 956–960. 11. Yaksi, E., and Wilson, R.I. (2010). Electrical coupling between olfactory glomeruli. Neuron 67, 1034–1047. 12. Huang, J., Zhang, W., Qiao, W., Hu, A., and Wang, Z. (2010). Functional connectivity and selective odor responses of excitatory local interneurons in Drosophila antennal lobe. Neuron 67, 1021–1033. 13. Olsen, S.R., Bhandawat, V., and Wilson, R.I. (2007). Excitatory interactions between olfactory processing channels in the Drosophila antennal lobe. Neuron 54, 89–103. 14. Kosaka, T., and Kosaka, K. (2003). Neuronal gap junctions in the rat main olfactory bulb, with special reference to intraglomerular gap junctions. Neurosci. Res. 45, 189–209. 15. Paternostro, M.A., Reyher, C.K., and Brunjes, P.C. (1995). Intracellular injections of lucifer yellow into lightly fixed mitral cells reveal neuronal dye-coupling in the developing rat olfactory bulb. Dev. Brain Res. 84, 1–10. 16. Shipley, M.T., and Ennis, M. (1996). Functional organization of olfactory system. J. Neurobiol. 30, 123–176. 17. Root, C.M., Masuyama, K., Green, D.S., Enell, L.E., Nassel, D.R., Lee, C.H., and Wang, J.W. (2008). A presynaptic gain control mechanism fine-tunes olfactory behavior. Neuron 59, 311–321. 18. Yoshihara, Y. (2009). Molecular genetic dissection of the zebrafish olfactory system. In Chemosensory Systems in Mammals, Fishes, and Insects, W. Meyerhof and S.I. Korsching, eds. (Heidelberg Berlin: Springer-Verlag), pp. 97–120. 19. Fuller, C.L., Yettaw, H.K., and Byrd, C.A. (2006). Mitral cells in the olfactory bulb of adult zebrafish (Danio rerio): morphology and distribution. J. Comp. Neurol. 499, 218–230. 20. Wilson, R.I. (2013). Early olfactory processing in Drosophila: mechanisms and principles. Annu. Rev. Neurosci. 36, 217–241.
Department of Neurobiology, Harvard Medical School, 220 Longwood Ave., Boston, MA 02115, USA. E-mail: [email protected], [email protected] http://dx.doi.org/10.1016/j.cub.2013.10.056
Evolution: Unveiling Early Alveolates The isolation and characterisation of a novel protist lineage enables the reconstruction of early evolutionary events that gave rise to ciliates, malaria parasites, and coral symbionts. These events include dramatic changes in mitochondrial genome content and organisation. Richard G. Dorrell1, Erin R. Butterfield1,2, R. Ellen R. Nisbet1,2, and Christopher J. Howe1,* Animals, plants, and other multicellular organisms are a drop in the ocean of eukaryotic diversity. A vast array of different protist lineages are known, many of which have
extremely important functions in planetary ecology, or are major human pathogens [1]. Some protists have served as models for processes of broad biological significance; for example, early work on telomere maintenance used the ciliate Tetrahymena [2]. However, compared to animals and other model eukaryotes, most of the cell biology of
Current Biology Vol 23 No 24 R1094
Dinoflagellates (coral symbionts, free-living and “red tide” algae) Hematodinium Apicomplexans (Plasmodium, Toxoplasma)
Ciliates (Paramecium, Tetrahymena)
Chromera B Colponema sp. Peru
Colponemids?
Colponemids?
A
Colponema edaphicum/ sp. Vietnam
Stramenopiles (kelps, diatoms, oomycetes)
Other eukaryotes Current Biology
Figure 1. Evolutionary relationships of colponemids to other lineages. This tree shows the diversity of colponemids, as described in [6], in relation to other key alveolate lineages. Dotted lines refer to colponemid lineages identified only from environmental sequences. Two key events in the evolution of the alveolate mitochondrial genome are shown in green: (A) the adoption of a linear genome form by an early alveolate that is ancestral to all currently studied lineages, following their divergence from their closest relatives, the stramenopiles, and (B) the massive reduction in genome content in a common ancestor of the apicomplexans and dinoflagellates, following their divergence from Colponema sp. Peru.
protists is extremely underexplored, partially due to the fact that very many species remain extremely difficult to establish in laboratory culture. Certain lineages that are believed to be highly diverse, or occupy important positions in the tree of eukaryotes, are known only from environmental data, or have only recently been established in culture [3–5]. Here, as reported in this issue of Current Biology, a new study kovec et al. [6] shows by Janous how a previously poorly understood eukaryotic
genus — Colponema — may provide major insights into the evolution of one of the most enigmatic groups, the alveolates. The alveolates are an ancient group of eukaryotes that occupy a diverse array of ecological niches, both free-living and parasitic. Alveolates form a sister-group to the stramenopiles, which include diatom algae, kelps, and oomycete pathogens, but are much more distant relatives of plants, animals and fungi [1]. Some of the most recognisable
members of the alveolates are the ciliates, a group of heterotrophic protists that include Tetrahymena and Paramecium, and play important roles in microbial food webs. The majority of ciliates are free-living predators, although parasitism appears to have evolved several times [1,7]. The dinoflagellates are another major alveolate group. They contain both heterotrophic and photosynthetic members, many of which form symbiotic associations, such as the photosynthetic ‘zooxanthellae’ of coral. In addition, there are the apicomplexans, a largely parasitic lineage, including the major pathogens Plasmodium (the causative agent of malaria) and Toxoplasma (toxoplasmosis) [1]. Ciliates diverge at the base of the alveolates, with the dinoflagellates and apicomplexans forming a single clade (Figure 1) [1,8]. In addition to their extraordinary diversity in life strategy, specific alveolate groups have unusual and highly distinctive forms of genome organisation. The ciliate nuclear genome is contained in two different organelles, with different forms: a vegetative ‘micronucleus’ that contains large numbers of gene fragments, and a ‘macronucleus’, containing translationally functional genes, which is generated via the rearrangement of the micronuclear genome [9]. Dinoflagellates have a permanently condensed nuclear genome, containing large tandem arrays of intron-rich genes, while apicomplexans have a much more conventional nuclear genome [10]. Ciliates lack chloroplasts, while dinoflagellates and apicomplexans retain highly unusual plastids. The dinoflagellate chloroplast genome is highly reduced in content, and fragmented into a number of small, plasmid-like elements termed ‘minicircles’. The apicomplexan chloroplast has lost all genes of photosynthetic function, and has been converted into a non-photosynthetic organelle termed the ‘apicoplast’ [8], which is an attractive target for drugs to combat diseases such as malaria. Unlike the situation in stramenopiles, or in other major eukaryotic groups, alveolate mitochondrial genomes are linear. However, while the mitochondrial genome of the free-living ciliates is rich in genes, the genomes of both dinoflagellates and apicomplexan
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mitochondria are highly reduced in content, typically encoding only three proteins and ribosomal RNA [6,11]. Understanding how and when the different alveolate genome organisation patterns originated may provide insights into their evolutionary history and, in the case of the apicomplexans, suggest targets for therapeutic treatment. However, reconstructing the evolutionary history of the alveolates is problematic, due to the deep divergence dates between ciliates, dinoflagellates and apicomplexans. It is believed that ciliates branched from other alveolates up to one billion years ago, while apicomplexans and dinoflagellates diverged more than three hundred million years before the present [12,13]. The recent characterisation of other alveolates that branch closer to the dinoflagellates (Hematodinium) and to the apicomplexans (Chromera) has provided valuable insights into the origins of the permanently condensed dinoflagellate nuclear genome, and the gene loss events that have given rise to the apicoplast [8,14]. Understanding the evolution of the highly reduced mitochondrial genomes of dinoflagellates and apicomplexans has been much more problematic, because of the lack of appropriate close relatives with which to compare them. In the most recent study reported kovec et al. in this issue, Janous demonstrate that Colponema, previously believed to be a single genus of free-living heterotrophs of uncertain phylogenetic placement, forms two phylogenetically distinct groups, which both resolve at important positions within the alveolates [6]. The previously described Colponema edaphicum and Colponema sp. Vietnam, a novel lineage established from environmental samples, group together at the base of studied alveolates (Figure 1). A third isolate, Colponema sp. Peru, forms a separate sister-group to apicomplexans and dinoflagellates (Figure 1), and accordingly provides a model for exploring mitochondrial evolution in these lineages. kovec et al. also present a Janous complete, linear, mitochondrial genome for the Peru isolate [6]. Surprisingly, this mitochondrial genome is extraordinarily gene-rich, containing ten times as many genes as those of apicomplexans or
dinoflagellates [6], and even retains genes that have been lost from ciliate mitochondria. The discovery of a species with an extremely rich mitochondrial genome that is closely related to lineages that have undergone an extraordinary degree of mitochondrial genome reduction raises important questions. Clearly, the genome reduction observed in certain alveolates must have occurred over the relatively short evolutionary time-period between the divergence of Colponema and the radiation of dinoflagellates and apicomplexans, so what enabled it to kovec et al. suggest occur? Janous that much of the mitochondrial gene loss in apicomplexans and dinoflagellates must have been dependent on the recruitment of cytoplasmic pathways to support the mitochondria [6]. The evolution of a tRNA import pathway for mitochondria, similar to the import machinery found in kinetoplastid parasites, could explain the loss of kovec tRNA genes [15,16]. Janous et al. also suggest loss of complex I NADH dehydrogenase genes may be associated with the substitution of an alternative type 2 NADH dehydrogenase, as is found in Plasmodium and Cryptosporidium. However, the situation is likely to be more complex than a direct substitution, as dinoflagellates may have both the conventional and alternative NADH dehydrogenases [13,17,18]. Another intriguing feature is the retention in Colponema sp. Peru of the atp6 gene, encoding the a subunit of the Fo component of the ATP synthase. atp6 has been lost in dinoflagellates, apicomplexans and ciliates, although Tetrahymena has been reported to possess a replacement gene [19,20]. Given the apparent phylogenetic position of Colponema sp. Peru between the ciliates and apicomplexans (and dinoflagellates), this suggests multiple independent changes to this important complex. These changes are unlikely to explain all of the gene loss events that occurred, and other evolutionary factors are probably also at play. While ciliates and Colponema have large mitochondrial genomes, and are believed to be generally free-living, both apicomplexans and dinoflagellates are frequently found in close association with other
organisms, respectively as parasites, and as symbionts and commensals. Perhaps a reduction in the apicomplexan mitochondrial genome might be associated with the transition from a free-living lifestyle strategy to one at least primed for the transition to parasitism. Increased sampling of mitochondrial genomes in colponemids and related species may provide an answer. Ultimately, the phylogenetic and genomic characterisation of Colponema is a major development in understanding the evolution of the alveolates. However, important questions remain. The much slower evolutionary rates associated with colponemid mitochondrial genes compared to those of ciliates, dinoflagellates, and apicomplexans is intriguing, and warrants further kovec et al. also investigation. Janous provide some striking preliminary data suggesting that other colponemid lineages, currently known only from environmental sequences, form independent branches on the tree of alveolates (Figure 1) [6]. Isolation and exploration of some of these lineages could provide much greater resolution of the early evolutionary events in alveolate history. As ever, currently uncultured protists have the potential to transform dramatically our understanding of the eukaryotic tree. References 1. Walker, G., Dorrell, R.G., Schlacht, A., and Dacks, J.B. (2011). Eukaryotic systematics: a user’s guide for cell biologists and parasitologists. Parasitol. 138, 1638–1663. 2. Greider, C.W., and Blackburn, E.H. (1985). Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 43, 405–413. 3. Kim, E., Harrison, J.W., Sudek, S., Jones, M.D., Wilcox, H.M., Richards, T.A., Worden, A.Z., and Archibald, J.M. (2011). Newly identified and diverse plastid-bearing branch on the eukaryotic tree of life. Proc. Natl. Acad. Sci. USA 108, 1496–1500. 4. Seenivasan, R., Sausen, N., Medlin, L.K., and Melkonian, M. (2013). Picomonas judraskeda gen. et sp. nov.: the first identified member of the Picozoa phylum nov., a widespread group of picoeukaryotes, formerly known as ‘picobiliphytes’. PLoS One 8, e59565. 5. Jones, M.D., Forn, I., Gadelha, C., Egan, M.J., Bass, D., Massana, R., and Richards, T.A. (2011). Discovery of novel intermediate forms redefines the fungal tree of life. Nature 474, 200–203. kovec, J., Tikhonenkov, D., 6. Janous Mikhailov, K., Simdyanov, T., Aleoshin, V., Mylnikov, A., and Keeling, P. (2013). Colponemids represent multiple ancient alveolate lineages. Curr. Biol. 23, 2546–2552. 7. Baker, J.R. (1994). The origins of parasitism in the protists. Int. J. Parasitol. 24, 1131–1137.
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kovec, J., Hora´k, A., Obornı´k, M., 8. Janous Lukes, J., and Keeling, P.J. (2010). A common red algal origin of the apicomplexan, dinoflagellate, and heterokont plastids. Proc. Natl. Acad. Sci. USA 107, 10949–10954. 9. Eisen, J.A., Coyne, R.S., Wu, M., Wu, D.Y., Thiagarajan, M., Wortman, J.R., Badger, J.H., Ren, Q.H., Amedeo, P., Jones, K.M., et al. (2006). Macronuclear genome sequence of the ciliate Tetrahymena thermophila, a model eukaryote. PLoS Biol. 4, 1620–1642. 10. Shoguchi, E., Shinzato, C., Kawashima, T., Gyoja, F., Mungpakdee, S., Koyanagi, R., Takeuchi, T., Hisata, K., Tanaka, M., Fujiwara, M., et al. (2013). Draft assembly of the Symbiodinium minutum nuclear genome reveals dinoflagellate gene structure. Curr. Biol. 23, 1399–1408. 11. Nash, E.A., Nisbet, R.E.R., Barbrook, A.C., and Howe, C.J. (2008). Dinoflagellates: a mitochondrial genome all at sea. Trends Genet. 24, 328–335. 12. Parfrey, L.W., Lahr, D.J., Knoll, A.H., and Katz, L.A. (2011). Estimating the timing of early eukaryotic diversification with multigene molecular clocks. Proc. Natl. Acad. Sci. USA 108, 13624–13629.
13. Butterfield, E.R., Howe, C.J., and Nisbet, R.E.R. (2013). An analysis of dinoflagellate metabolism using EST data. Protist 164, 218–236. 14. Gornik, S.G., Ford, K.L., Mulhern, T.D., Bacic, A., McFadden, G.I., and Waller, R.F. (2012). Loss of nucleosomal DNA condensation coincides with appearance of a novel nuclear protein in dinoflagellates. Curr. Biol. 22, 2303–2312. 15. Seidman, D., Johnson, D., Gerbasi, V., Golden, D., Orlando, R., and Hajduk, S. (2012). Mitochondrial membrane complex that contains proteins necessary for tRNA import in Trypanosoma brucei. J. Biol. Chem. 287, 8892–8903. 16. Esseiva, A.C., Naguleswaran, A., Hemphill, A., and Schneider, A. (2004). Mitochondrial tRNA import in Toxoplasma gondii. J. Biol. Chem. 279, 42363–42368. 17. Mogi, T., and Kita, K. (2010). Diversity in mitochondrial metabolic pathways in parasitic protists Plasmodium and Cryptosporidium. Parasitol. Int. 59, 305–312. 18. Kerscher, S.J. (2000). Diversity and origin of alternative NADH: ubiquinone oxidoreductases. Biochim. Biophys. Acta 1459, 274–283.
Early Vision: Where (Some of) the Magic Happens A recent study provides new insights into how the very different response characteristics of the main visual pathways from the eye to the brain may directly result from the presence or absence of a ‘spike trigger-zone’ in retinal bipolar cells. Tom Baden and Thomas Euler The visual system of primates, including that of humans, famously features both exquisite spatial acuity and a high temporal resolution. This dual focus on both ‘sharpness’ and ‘speed’ is made possible through different processing streams set up already in the retina. In a recent study, Puthussery et al. [1] now show that key differences in the processing streams that are thought to underlie these visual abilities are already set up right after the first synapse of the visual system — in retinal bipolar cells. Origin of Magnocellular and Parvocellular Channels The retina breaks the visual world into several parallel representations prior to transmission to the brain. Each representation, or ‘channel’, is based on a different type of retinal ganglion cell that carries information about specific features of the visual scene — such as edges, directed motion or ‘color’ [2]. Of the 20 or so types of ganglion cells that exist in the primate retina, two in particular
have attracted considerable attention since they were first described in the 1940s [3]: the parasol and midget cells. Parasol ganglion cells have large receptive fields, display transient visual responses and implement the ‘magnocellular’ (M) pathway. Midget ganglion cells, in contrast, have tiny receptive fields, slow responses and provide the ‘parvocellular’ (P) pathway (Figure 1A). Based on their physiology and projection targets in the brain, the magnocellular versus parvocellular channels have long been implicated as the source of high temporal precision and high spatial acuity in primate vision, respectively (for example [4]). But how are the striking differences in midget versus parasol cell light responses derived from their respective retinal microcircuits? Much of the distinct response signature of ganglion cells is determined by their complement of synaptic inputs, namely excitatory drive from bipolar cells, and inhibition provided by amacrine cells [5,6]. Midget ganglion cells are driven by midget bipolar cells, while parasol ganglion cells receive major inputs from so-called ‘DB3’ and
19. Balabaskaran Nina, P., Dudkina, N.V., Kane, L.A., van Eyk, J.E., Boekema, E.J., Mather, M.W., and Vaidya, A.B. (2010). Highly divergent mitochondrial ATP synthase complexes in Tetrahymena thermophila. PLoS Biol. 8, 1000418. 20. Nisbet, R.E.R., Mitton, E.R., and Menz, R.I. (2011). A highly divergent mitochondrial ATP synthase which confirms the monophyly of the chromalveolates. ICOPA Proc. 12, 105–108.
1Department
of Biochemistry, University of Cambridge, Building O, Downing Site, Tennis Court Road, Cambridge, CB2 1QW, UK. 2School of Pharmacy and Medical Sciences, University of South Australia, North Terrace, Adelaide, SA 5000, Australia. *E-mail: [email protected]
http://dx.doi.org/10.1016/j.cub.2013.10.055
‘DB4’ bipolar cells (DB referring to ‘diffuse bipolar’ cell). Hence, midget bipolar cells should exhibit a slow, sustained physiology, whereas DB3/4 cells should be more transient. To address this long standing question, Puthussery et al. [1] investigated what active conductances are present in primate DB3/4 bipolar cells and how the underlying ion channels are distributed. In their beautiful study, the authors show that a specific set of ion channels is present in DB3/4 bipolar cells but completely absent in midget bipolar cells. These channels, most notably including voltage-gated sodium channels, appear to work together to render DB3/ 4 cells intrinsically highly transient and nonlinear. The channel complement of DB3/4 cells even allows them to generate full-blown action potentials — very much unlike the midget bipolar cells, which behaved in a fully graded, almost ‘passive’ manner (Figure 1A). Pharmacological blockage of these sodium channels resulted in much less transient light-driven inputs to parasol, but not to midget ganglion cells. In this way, Puthussery et al. [1] build on a growing body of evidence that active conductances and spikes in bipolar cells present a fundamental ingredient for high-precision temporal processing in the visual system [7–13]. The Design of a Retinal Bipolar Cell The study by Puthussery et al. [1] goes beyond the ‘mere’ demonstration
Ultrastructural and dye filling studies have suggested that GABAergic interneurons make gap junctions onto mitral cells in the mammalian bulb [14,15]. However, this remains a controversial topic that requires further work. New Questions The study by Zhu et al. [1] raises many questions. First, exactly which interneurons are involved? The dlx4/6 neurons are a heterogeneous population [1], like the interneurons of the Drosophila antennal lobe, and some of these neurons may play entirely distinct roles. Identifying a genetic marker for each relevant interneuron type would permit more targeted recordings, better mapping of connectivity, and more precise manipulations. It is also notable that there are likely many other GABAergic interneurons in the fish olfactory bulb in addition to the dlx4/6 neurons. In the mammalian bulb there are several populations of periglomerular cells in the superficial glomerular layer, as well as deep-layer granule cells, which outnumber all other cell types in the bulb by at least an order of magnitude [16]. What specific roles do each of these different inhibitory circuits play, and how do they interact? Second, do specific odor stimuli elicit specific spatial patterns of interactions among olfactory glomeruli? Or alternatively, is this a global interaction that simply scales in strength with the total level of afferent input to the circuit? None of the studies discussed here provides a clear answer to this question. This issue is critical to understanding how interneurons affect olfactory processing. Third, why does this circuit have such diverse effects on different target neurons? When the dlx4/6 neurons were manipulated, there were large variations across cells in the effects this had on neural activity. For a given odor stimulus, some cells were inhibited, but others were disinhibited [1]. Again, this finding has a parallel in the Drosophila antennal lobe, where identified olfactory glomeruli have diverse levels of sensitivity to lateral inhibition and lateral excitation [8,13,17]. What are the mechanisms and significance of this diversity? Finally, how deep are the functional and structural similarities between neural circuits in different organisms? We should remember that the success
of molecular biology in the 20th century hinged on our ability to spot homologies between amino acid sequences in different protein domains, and to grasp the systematic relationship between the primary structure of a domain and its function. Many such molecular modules are now known to reoccur throughout the animal kingdom. Being able to spot similar kinds of structure–function relationships in neural circuits would accelerate discovery in neuroscience. For this reason, comparative biology remains essential to the search for general principles. References 1. Zhu, P., Frank, T., and Friedrich, R.W. (2013). Equalization of odor representations by a network of electrically coupled inhibitory interneurons. Nat Neurosci. 16, 1678–1686. 2. Fairhall, A. (2014). Adaptation and natural stimulus statistics. In The Cognitive Neurosciences 5th Edition, M.S. Gazzaniga, ed. (Cambridge, MA: MIT Press). 3. Laughlin, S.B. (2001). Energy as a constraint on the coding and processing of sensory information. Curr. Opin. Neurobiol. 11, 475–480. 4. Shapley, R., and Enroth-Cugell, C. (1984). Visual adaptation and retinal gain controls. Prog. Retin. Res. 3, 263–346. 5. Carandini, M., and Heeger, D.J. (2012). Normalization as a canonical neural computation. Nat. Rev. Neurosci. 13, 51–62. 6. Zoccolan, D., Cox, D.D., and DiCarlo, J.J. (2005). Multiple object response normalization in monkey inferotemporal cortex. J. Neurosci. 25, 8150–8164. 7. Friedrich, R.W., and Korsching, S.I. (1997). Combinatorial and chemotopic odorant coding in the zebrafish olfactory bulb visualized by optical imaging. Neuron 18, 737–752. 8. Olsen, S.R., Bhandawat, V., and Wilson, R.I. (2010). Divisive normalization in olfactory population codes. Neuron 66, 287–299. 9. Asahina, K., Louis, M., Piccinotti, S., and Vosshall, L.B. (2009). A circuit supporting concentration-invariant odor perception in Drosophila. J. Biol. 8, 9.
10. Olsen, S.R., and Wilson, R.I. (2008). Lateral presynaptic inhibition mediates gain control in an olfactory circuit. Nature 452, 956–960. 11. Yaksi, E., and Wilson, R.I. (2010). Electrical coupling between olfactory glomeruli. Neuron 67, 1034–1047. 12. Huang, J., Zhang, W., Qiao, W., Hu, A., and Wang, Z. (2010). Functional connectivity and selective odor responses of excitatory local interneurons in Drosophila antennal lobe. Neuron 67, 1021–1033. 13. Olsen, S.R., Bhandawat, V., and Wilson, R.I. (2007). Excitatory interactions between olfactory processing channels in the Drosophila antennal lobe. Neuron 54, 89–103. 14. Kosaka, T., and Kosaka, K. (2003). Neuronal gap junctions in the rat main olfactory bulb, with special reference to intraglomerular gap junctions. Neurosci. Res. 45, 189–209. 15. Paternostro, M.A., Reyher, C.K., and Brunjes, P.C. (1995). Intracellular injections of lucifer yellow into lightly fixed mitral cells reveal neuronal dye-coupling in the developing rat olfactory bulb. Dev. Brain Res. 84, 1–10. 16. Shipley, M.T., and Ennis, M. (1996). Functional organization of olfactory system. J. Neurobiol. 30, 123–176. 17. Root, C.M., Masuyama, K., Green, D.S., Enell, L.E., Nassel, D.R., Lee, C.H., and Wang, J.W. (2008). A presynaptic gain control mechanism fine-tunes olfactory behavior. Neuron 59, 311–321. 18. Yoshihara, Y. (2009). Molecular genetic dissection of the zebrafish olfactory system. In Chemosensory Systems in Mammals, Fishes, and Insects, W. Meyerhof and S.I. Korsching, eds. (Heidelberg Berlin: Springer-Verlag), pp. 97–120. 19. Fuller, C.L., Yettaw, H.K., and Byrd, C.A. (2006). Mitral cells in the olfactory bulb of adult zebrafish (Danio rerio): morphology and distribution. J. Comp. Neurol. 499, 218–230. 20. Wilson, R.I. (2013). Early olfactory processing in Drosophila: mechanisms and principles. Annu. Rev. Neurosci. 36, 217–241.
Department of Neurobiology, Harvard Medical School, 220 Longwood Ave., Boston, MA 02115, USA. E-mail: [email protected], [email protected] http://dx.doi.org/10.1016/j.cub.2013.10.056
Evolution: Unveiling Early Alveolates The isolation and characterisation of a novel protist lineage enables the reconstruction of early evolutionary events that gave rise to ciliates, malaria parasites, and coral symbionts. These events include dramatic changes in mitochondrial genome content and organisation. Richard G. Dorrell1, Erin R. Butterfield1,2, R. Ellen R. Nisbet1,2, and Christopher J. Howe1,* Animals, plants, and other multicellular organisms are a drop in the ocean of eukaryotic diversity. A vast array of different protist lineages are known, many of which have
extremely important functions in planetary ecology, or are major human pathogens [1]. Some protists have served as models for processes of broad biological significance; for example, early work on telomere maintenance used the ciliate Tetrahymena [2]. However, compared to animals and other model eukaryotes, most of the cell biology of
Current Biology Vol 23 No 24 R1094
Dinoflagellates (coral symbionts, free-living and “red tide” algae) Hematodinium Apicomplexans (Plasmodium, Toxoplasma)
Ciliates (Paramecium, Tetrahymena)
Chromera B Colponema sp. Peru
Colponemids?
Colponemids?
A
Colponema edaphicum/ sp. Vietnam
Stramenopiles (kelps, diatoms, oomycetes)
Other eukaryotes Current Biology
Figure 1. Evolutionary relationships of colponemids to other lineages. This tree shows the diversity of colponemids, as described in [6], in relation to other key alveolate lineages. Dotted lines refer to colponemid lineages identified only from environmental sequences. Two key events in the evolution of the alveolate mitochondrial genome are shown in green: (A) the adoption of a linear genome form by an early alveolate that is ancestral to all currently studied lineages, following their divergence from their closest relatives, the stramenopiles, and (B) the massive reduction in genome content in a common ancestor of the apicomplexans and dinoflagellates, following their divergence from Colponema sp. Peru.
protists is extremely underexplored, partially due to the fact that very many species remain extremely difficult to establish in laboratory culture. Certain lineages that are believed to be highly diverse, or occupy important positions in the tree of eukaryotes, are known only from environmental data, or have only recently been established in culture [3–5]. Here, as reported in this issue of Current Biology, a new study kovec et al. [6] shows by Janous how a previously poorly understood eukaryotic
genus — Colponema — may provide major insights into the evolution of one of the most enigmatic groups, the alveolates. The alveolates are an ancient group of eukaryotes that occupy a diverse array of ecological niches, both free-living and parasitic. Alveolates form a sister-group to the stramenopiles, which include diatom algae, kelps, and oomycete pathogens, but are much more distant relatives of plants, animals and fungi [1]. Some of the most recognisable
members of the alveolates are the ciliates, a group of heterotrophic protists that include Tetrahymena and Paramecium, and play important roles in microbial food webs. The majority of ciliates are free-living predators, although parasitism appears to have evolved several times [1,7]. The dinoflagellates are another major alveolate group. They contain both heterotrophic and photosynthetic members, many of which form symbiotic associations, such as the photosynthetic ‘zooxanthellae’ of coral. In addition, there are the apicomplexans, a largely parasitic lineage, including the major pathogens Plasmodium (the causative agent of malaria) and Toxoplasma (toxoplasmosis) [1]. Ciliates diverge at the base of the alveolates, with the dinoflagellates and apicomplexans forming a single clade (Figure 1) [1,8]. In addition to their extraordinary diversity in life strategy, specific alveolate groups have unusual and highly distinctive forms of genome organisation. The ciliate nuclear genome is contained in two different organelles, with different forms: a vegetative ‘micronucleus’ that contains large numbers of gene fragments, and a ‘macronucleus’, containing translationally functional genes, which is generated via the rearrangement of the micronuclear genome [9]. Dinoflagellates have a permanently condensed nuclear genome, containing large tandem arrays of intron-rich genes, while apicomplexans have a much more conventional nuclear genome [10]. Ciliates lack chloroplasts, while dinoflagellates and apicomplexans retain highly unusual plastids. The dinoflagellate chloroplast genome is highly reduced in content, and fragmented into a number of small, plasmid-like elements termed ‘minicircles’. The apicomplexan chloroplast has lost all genes of photosynthetic function, and has been converted into a non-photosynthetic organelle termed the ‘apicoplast’ [8], which is an attractive target for drugs to combat diseases such as malaria. Unlike the situation in stramenopiles, or in other major eukaryotic groups, alveolate mitochondrial genomes are linear. However, while the mitochondrial genome of the free-living ciliates is rich in genes, the genomes of both dinoflagellates and apicomplexan
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mitochondria are highly reduced in content, typically encoding only three proteins and ribosomal RNA [6,11]. Understanding how and when the different alveolate genome organisation patterns originated may provide insights into their evolutionary history and, in the case of the apicomplexans, suggest targets for therapeutic treatment. However, reconstructing the evolutionary history of the alveolates is problematic, due to the deep divergence dates between ciliates, dinoflagellates and apicomplexans. It is believed that ciliates branched from other alveolates up to one billion years ago, while apicomplexans and dinoflagellates diverged more than three hundred million years before the present [12,13]. The recent characterisation of other alveolates that branch closer to the dinoflagellates (Hematodinium) and to the apicomplexans (Chromera) has provided valuable insights into the origins of the permanently condensed dinoflagellate nuclear genome, and the gene loss events that have given rise to the apicoplast [8,14]. Understanding the evolution of the highly reduced mitochondrial genomes of dinoflagellates and apicomplexans has been much more problematic, because of the lack of appropriate close relatives with which to compare them. In the most recent study reported kovec et al. in this issue, Janous demonstrate that Colponema, previously believed to be a single genus of free-living heterotrophs of uncertain phylogenetic placement, forms two phylogenetically distinct groups, which both resolve at important positions within the alveolates [6]. The previously described Colponema edaphicum and Colponema sp. Vietnam, a novel lineage established from environmental samples, group together at the base of studied alveolates (Figure 1). A third isolate, Colponema sp. Peru, forms a separate sister-group to apicomplexans and dinoflagellates (Figure 1), and accordingly provides a model for exploring mitochondrial evolution in these lineages. kovec et al. also present a Janous complete, linear, mitochondrial genome for the Peru isolate [6]. Surprisingly, this mitochondrial genome is extraordinarily gene-rich, containing ten times as many genes as those of apicomplexans or
dinoflagellates [6], and even retains genes that have been lost from ciliate mitochondria. The discovery of a species with an extremely rich mitochondrial genome that is closely related to lineages that have undergone an extraordinary degree of mitochondrial genome reduction raises important questions. Clearly, the genome reduction observed in certain alveolates must have occurred over the relatively short evolutionary time-period between the divergence of Colponema and the radiation of dinoflagellates and apicomplexans, so what enabled it to kovec et al. suggest occur? Janous that much of the mitochondrial gene loss in apicomplexans and dinoflagellates must have been dependent on the recruitment of cytoplasmic pathways to support the mitochondria [6]. The evolution of a tRNA import pathway for mitochondria, similar to the import machinery found in kinetoplastid parasites, could explain the loss of kovec tRNA genes [15,16]. Janous et al. also suggest loss of complex I NADH dehydrogenase genes may be associated with the substitution of an alternative type 2 NADH dehydrogenase, as is found in Plasmodium and Cryptosporidium. However, the situation is likely to be more complex than a direct substitution, as dinoflagellates may have both the conventional and alternative NADH dehydrogenases [13,17,18]. Another intriguing feature is the retention in Colponema sp. Peru of the atp6 gene, encoding the a subunit of the Fo component of the ATP synthase. atp6 has been lost in dinoflagellates, apicomplexans and ciliates, although Tetrahymena has been reported to possess a replacement gene [19,20]. Given the apparent phylogenetic position of Colponema sp. Peru between the ciliates and apicomplexans (and dinoflagellates), this suggests multiple independent changes to this important complex. These changes are unlikely to explain all of the gene loss events that occurred, and other evolutionary factors are probably also at play. While ciliates and Colponema have large mitochondrial genomes, and are believed to be generally free-living, both apicomplexans and dinoflagellates are frequently found in close association with other
organisms, respectively as parasites, and as symbionts and commensals. Perhaps a reduction in the apicomplexan mitochondrial genome might be associated with the transition from a free-living lifestyle strategy to one at least primed for the transition to parasitism. Increased sampling of mitochondrial genomes in colponemids and related species may provide an answer. Ultimately, the phylogenetic and genomic characterisation of Colponema is a major development in understanding the evolution of the alveolates. However, important questions remain. The much slower evolutionary rates associated with colponemid mitochondrial genes compared to those of ciliates, dinoflagellates, and apicomplexans is intriguing, and warrants further kovec et al. also investigation. Janous provide some striking preliminary data suggesting that other colponemid lineages, currently known only from environmental sequences, form independent branches on the tree of alveolates (Figure 1) [6]. Isolation and exploration of some of these lineages could provide much greater resolution of the early evolutionary events in alveolate history. As ever, currently uncultured protists have the potential to transform dramatically our understanding of the eukaryotic tree. References 1. Walker, G., Dorrell, R.G., Schlacht, A., and Dacks, J.B. (2011). Eukaryotic systematics: a user’s guide for cell biologists and parasitologists. Parasitol. 138, 1638–1663. 2. Greider, C.W., and Blackburn, E.H. (1985). Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 43, 405–413. 3. Kim, E., Harrison, J.W., Sudek, S., Jones, M.D., Wilcox, H.M., Richards, T.A., Worden, A.Z., and Archibald, J.M. (2011). Newly identified and diverse plastid-bearing branch on the eukaryotic tree of life. Proc. Natl. Acad. Sci. USA 108, 1496–1500. 4. Seenivasan, R., Sausen, N., Medlin, L.K., and Melkonian, M. (2013). Picomonas judraskeda gen. et sp. nov.: the first identified member of the Picozoa phylum nov., a widespread group of picoeukaryotes, formerly known as ‘picobiliphytes’. PLoS One 8, e59565. 5. Jones, M.D., Forn, I., Gadelha, C., Egan, M.J., Bass, D., Massana, R., and Richards, T.A. (2011). Discovery of novel intermediate forms redefines the fungal tree of life. Nature 474, 200–203. kovec, J., Tikhonenkov, D., 6. Janous Mikhailov, K., Simdyanov, T., Aleoshin, V., Mylnikov, A., and Keeling, P. (2013). Colponemids represent multiple ancient alveolate lineages. Curr. Biol. 23, 2546–2552. 7. Baker, J.R. (1994). The origins of parasitism in the protists. Int. J. Parasitol. 24, 1131–1137.
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kovec, J., Hora´k, A., Obornı´k, M., 8. Janous Lukes, J., and Keeling, P.J. (2010). A common red algal origin of the apicomplexan, dinoflagellate, and heterokont plastids. Proc. Natl. Acad. Sci. USA 107, 10949–10954. 9. Eisen, J.A., Coyne, R.S., Wu, M., Wu, D.Y., Thiagarajan, M., Wortman, J.R., Badger, J.H., Ren, Q.H., Amedeo, P., Jones, K.M., et al. (2006). Macronuclear genome sequence of the ciliate Tetrahymena thermophila, a model eukaryote. PLoS Biol. 4, 1620–1642. 10. Shoguchi, E., Shinzato, C., Kawashima, T., Gyoja, F., Mungpakdee, S., Koyanagi, R., Takeuchi, T., Hisata, K., Tanaka, M., Fujiwara, M., et al. (2013). Draft assembly of the Symbiodinium minutum nuclear genome reveals dinoflagellate gene structure. Curr. Biol. 23, 1399–1408. 11. Nash, E.A., Nisbet, R.E.R., Barbrook, A.C., and Howe, C.J. (2008). Dinoflagellates: a mitochondrial genome all at sea. Trends Genet. 24, 328–335. 12. Parfrey, L.W., Lahr, D.J., Knoll, A.H., and Katz, L.A. (2011). Estimating the timing of early eukaryotic diversification with multigene molecular clocks. Proc. Natl. Acad. Sci. USA 108, 13624–13629.
13. Butterfield, E.R., Howe, C.J., and Nisbet, R.E.R. (2013). An analysis of dinoflagellate metabolism using EST data. Protist 164, 218–236. 14. Gornik, S.G., Ford, K.L., Mulhern, T.D., Bacic, A., McFadden, G.I., and Waller, R.F. (2012). Loss of nucleosomal DNA condensation coincides with appearance of a novel nuclear protein in dinoflagellates. Curr. Biol. 22, 2303–2312. 15. Seidman, D., Johnson, D., Gerbasi, V., Golden, D., Orlando, R., and Hajduk, S. (2012). Mitochondrial membrane complex that contains proteins necessary for tRNA import in Trypanosoma brucei. J. Biol. Chem. 287, 8892–8903. 16. Esseiva, A.C., Naguleswaran, A., Hemphill, A., and Schneider, A. (2004). Mitochondrial tRNA import in Toxoplasma gondii. J. Biol. Chem. 279, 42363–42368. 17. Mogi, T., and Kita, K. (2010). Diversity in mitochondrial metabolic pathways in parasitic protists Plasmodium and Cryptosporidium. Parasitol. Int. 59, 305–312. 18. Kerscher, S.J. (2000). Diversity and origin of alternative NADH: ubiquinone oxidoreductases. Biochim. Biophys. Acta 1459, 274–283.
Early Vision: Where (Some of) the Magic Happens A recent study provides new insights into how the very different response characteristics of the main visual pathways from the eye to the brain may directly result from the presence or absence of a ‘spike trigger-zone’ in retinal bipolar cells. Tom Baden and Thomas Euler The visual system of primates, including that of humans, famously features both exquisite spatial acuity and a high temporal resolution. This dual focus on both ‘sharpness’ and ‘speed’ is made possible through different processing streams set up already in the retina. In a recent study, Puthussery et al. [1] now show that key differences in the processing streams that are thought to underlie these visual abilities are already set up right after the first synapse of the visual system — in retinal bipolar cells. Origin of Magnocellular and Parvocellular Channels The retina breaks the visual world into several parallel representations prior to transmission to the brain. Each representation, or ‘channel’, is based on a different type of retinal ganglion cell that carries information about specific features of the visual scene — such as edges, directed motion or ‘color’ [2]. Of the 20 or so types of ganglion cells that exist in the primate retina, two in particular
have attracted considerable attention since they were first described in the 1940s [3]: the parasol and midget cells. Parasol ganglion cells have large receptive fields, display transient visual responses and implement the ‘magnocellular’ (M) pathway. Midget ganglion cells, in contrast, have tiny receptive fields, slow responses and provide the ‘parvocellular’ (P) pathway (Figure 1A). Based on their physiology and projection targets in the brain, the magnocellular versus parvocellular channels have long been implicated as the source of high temporal precision and high spatial acuity in primate vision, respectively (for example [4]). But how are the striking differences in midget versus parasol cell light responses derived from their respective retinal microcircuits? Much of the distinct response signature of ganglion cells is determined by their complement of synaptic inputs, namely excitatory drive from bipolar cells, and inhibition provided by amacrine cells [5,6]. Midget ganglion cells are driven by midget bipolar cells, while parasol ganglion cells receive major inputs from so-called ‘DB3’ and
19. Balabaskaran Nina, P., Dudkina, N.V., Kane, L.A., van Eyk, J.E., Boekema, E.J., Mather, M.W., and Vaidya, A.B. (2010). Highly divergent mitochondrial ATP synthase complexes in Tetrahymena thermophila. PLoS Biol. 8, 1000418. 20. Nisbet, R.E.R., Mitton, E.R., and Menz, R.I. (2011). A highly divergent mitochondrial ATP synthase which confirms the monophyly of the chromalveolates. ICOPA Proc. 12, 105–108.
1Department
of Biochemistry, University of Cambridge, Building O, Downing Site, Tennis Court Road, Cambridge, CB2 1QW, UK. 2School of Pharmacy and Medical Sciences, University of South Australia, North Terrace, Adelaide, SA 5000, Australia. *E-mail: [email protected]
http://dx.doi.org/10.1016/j.cub.2013.10.055
‘DB4’ bipolar cells (DB referring to ‘diffuse bipolar’ cell). Hence, midget bipolar cells should exhibit a slow, sustained physiology, whereas DB3/4 cells should be more transient. To address this long standing question, Puthussery et al. [1] investigated what active conductances are present in primate DB3/4 bipolar cells and how the underlying ion channels are distributed. In their beautiful study, the authors show that a specific set of ion channels is present in DB3/4 bipolar cells but completely absent in midget bipolar cells. These channels, most notably including voltage-gated sodium channels, appear to work together to render DB3/ 4 cells intrinsically highly transient and nonlinear. The channel complement of DB3/4 cells even allows them to generate full-blown action potentials — very much unlike the midget bipolar cells, which behaved in a fully graded, almost ‘passive’ manner (Figure 1A). Pharmacological blockage of these sodium channels resulted in much less transient light-driven inputs to parasol, but not to midget ganglion cells. In this way, Puthussery et al. [1] build on a growing body of evidence that active conductances and spikes in bipolar cells present a fundamental ingredient for high-precision temporal processing in the visual system [7–13]. The Design of a Retinal Bipolar Cell The study by Puthussery et al. [1] goes beyond the ‘mere’ demonstration
