- Email: [email protected]
Real-time imaging reveals spatiotemporal dynamics of cellular circadian clocks
Update
TRENDS in Neurosciences Vol.27 No.9 September 2004
neuronal circuitry. Adult-fly-restricted expression of dominantly acting genes that inhibit evoked neurotransmitter release (e.g. UAS-tetanus toxin light chain [24]) or electrically silence neurons (e.g. UAS-Kir2.1, a genetically engineered KC leak channel [24]) will allow us to better understand the neural circuits underlying complex adult behaviors such as courtship, circadian rhythm, and learning and memory. Another powerful use of Gene-Switch and TARGET would be adult-fly-restricted RNA interference. UAS-driven double-stranded RNA hairpin transgenes [25] combined with TARGET or Gene-Switch could allow neurogeneticists to dissect the role of a gene in adult behavior independent of its role in neurodevelopment. The TARGET and Gene-Switch systems represent a significant technological advance for genetic studies of behavior in Drosophila. Prior to their development, memory researchers had to compromise, selecting either spatially restricted, semi-constitutive gene expression using the standard GAL4-UAS system or temporally controllable, spatially indiscriminate gene expression using the heatshock promoter. Thanks to the efforts of the Davis laboratory, these limitations are now merely memories.
References 1 McGuire, S.E. et al. (2003) Spatiotemporal rescue of memory dysfunction in Drosophila. Science 302, 1765–1768 2 Mao, Z. et al. (2004) Pharmacogenetic rescue in time and space of the rutabaga memory impairment by using Gene-Switch. Proc. Natl. Acad. Sci. U. S. A. 101, 198–203 3 Brand, A.H. and Perrimon, N. (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118, 401–415 4 Duffy, J.B. (2002) GAL4 system in Drosophila: a fly geneticist’s Swiss army knife. Genesis 34, 1–15 5 Lee, T. and Luo, L. (1999) Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22, 451–461 6 Burcin, M.M. et al. (1999) Adenovirus-mediated regulable target gene expression in vivo. Proc. Natl. Acad. Sci. U. S. A. 96, 355–360 7 Osterwalder, T. et al. (2001) A conditional tissue-specific transgene expression system using inducible GAL4. Proc. Natl. Acad. Sci. U. S. A. 98, 12596–12601
513
8 Roman, G. et al. (2001) P[Switch], a system for spatial and temporal control of gene expression in Drosophila melanogaster. Proc. Natl. Acad. Sci. U. S. A. 98, 12602–12607 9 Stebbins, M.J. et al. (2001) Tetracycline-inducible systems for Drosophila. Proc. Natl. Acad. Sci. U. S. A. 98, 10775–10780 10 Bello, B. et al. (1998) Spatial and temporal targeting of gene expression in Drosophila by means of a tetracycline-dependent transactivator system. Development 125, 2193–2202 11 Bieschke, E.T. et al. (1998) Doxycycline-induced transgene expression during Drosophila development and aging. Mol. Gen. Genet. 258, 571–579 12 Han, D.D. et al. (2000) Investigating the function of follicular subpopulations during Drosophila oogenesis through hormonedependent enhancer-targeted cell ablation. Development 127, 573–583 13 Levin, L.R. et al. (1992) The Drosophila learning and memory gene rutabaga encodes a Ca2C/calmodulin-responsive adenylyl cyclase. Cell 68, 479–489 14 Kandel, E.R. et al. (1983) Classical conditioning and sensitization share aspects of the same molecular cascade in Aplysia. Cold Spring Harb. Symp. Quant. Biol. 48, 821–830 15 Livingstone, M.S. et al. (1984) Loss of calcium/calmodulin responsiveness in adenylate cyclase of rutabaga, a Drosophila learning mutant. Cell 37, 205–215 16 Han, P.L. et al. (1992) Preferential expression of the Drosophila rutabaga gene in mushroom bodies, neural centers for learning in insects. Neuron 9, 619–627 17 Zars, T. et al. (2000) Localization of a short-term memory in Drosophila. Science 288, 672–675 18 Corfas, G. and Dudai, Y. (1991) Morphology of a sensory neuron in Drosophila is abnormal in memory mutants and changes during aging. Proc. Natl. Acad. Sci. U. S. A. 88, 7252–7256 19 Balling, A. et al. (1987) Are the structural changes in adult Drosophila mushroom bodies memory traces? Studies on biochemical learning mutants. J. Neurogenet. 4, 65–73 20 McCall, K. and Steller, H. (1997) Facing death in the fly: genetic analysis of apoptosis in Drosophila. Trends Genet. 13, 222–226 21 Roman, G. and Davis, R.L. (2002) Conditional expression of UAS-transgenes in the adult eye with a new gene-switch vector system. Genesis 34, 127–131 22 Rensing, L. and Ruoff, P. (2002) Temperature effect on entrainment, phase shifting, and amplitude of circadian clocks and its molecular bases. Chronobiol. Int. 19, 807–864 23 McGuire, S. et al. (2004) Spatiotemporal gene expression targeting with the TARGET and Gene-Switch systems in Drosophila. Sci. STKE 220, 14 24 White, B. et al. (2001) Molecular genetic approaches to the targeted suppression of neuronal activity. Curr. Biol. 11, R1041–R1053 25 Kalidas, S. and Smith, D.P. (2002) Novel genomic cDNA hybrids produce effective RNA interference in adult Drosophila. Neuron 33, 177–184 0166-2236/$ - see front matter Q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.tins.2004.06.013
Real-time imaging reveals spatiotemporal dynamics of cellular circadian clocks William J. Schwartz1 and Johanna H. Meijer2 1 2
Department of Neurology, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, MA 01655, USA Department of Neurophysiology, Leiden University Medical Centre, 2300 RC Leiden, The Netherlands
Circadian clocks have been localized to discrete sites within the nervous systems of several organisms, and in mammals to the suprachiasmatic nucleus (SCN) in the anterior hypothalamus. The clock in the SCN is Corresponding author: William J. Schwartz ([email protected]). Available online 10 July 2004 www.sciencedirect.com
composed of multiple autonomous single-cell oscillators, and new studies now allow an unprecedented look at their oscillatory activities over repeated cycles in tissue slices in vitro. Initial data reinforce the importance of intercellular membrane events for constructing a functional and reliable tissue pacemaker.
514
Update
TRENDS in Neurosciences Vol.27 No.9 September 2004
To adapt to the once-daily rotation of our planet about its axis, most organisms have evolved to incorporate the geophysical day–night cycle as a genetically determined temporal program. This endogenous, temperaturecompensated timekeeping mechanism – the ‘circadian clock’ – recognizes local time, measures its passage, and contributes to the regulation of homeostasis, seasonal behavior and navigation. Our progress in understanding its neurobiology has been almost unimaginable. About 30 years ago, the suprachiasmatic nucleus (SCN) was first implicated as the site of a master clock, but it has been less than ten years since SCN neurons were discovered to be autonomous cellular circadian oscillators [1]: when cultured at low density, they individually express firing rate rhythms with different circadian periods. Analyses of induced and spontaneous mutations, gene sequence homologies and protein–protein interactions have identified candidate regulatory molecules and biochemical processes that are likely to constitute the basic intracellular oscillatory mechanism. It is believed that genes at the core of the clock function within autoregulatory feedback loops, with nuclear proteins rhythmically suppressing the transcription of their own mRNA. Still mysterious is how individual SCN cells are assembled to create an integrated tissue pacemaker that governs circadian behaviors of whole animals. The genotype-specific circadian period characteristically expressed by animals appears to be an intermediate period that arises from the coupling of multiple SCN cellular oscillators [2–4]; this contrasts with many other rhythmic tissues (e.g. the heart), in which the fastest cells set the rate. Elucidating SCN intercellular interactions will be crucial to understanding circadian properties at the ‘tissue’ level of organization; several attributes of the clock are likely to emerge at this level, such as its regulation of complex behavioral patterns, adaptation to photoperiod [5] and remarkable precision [6]. Time for new technologies Recently there have been exciting advances in developing methods that make possible real-time simultaneous measurements of oscillatory activity over repeated cycles from multiple individual cells in tissue slices [7,8]. In a remarkable and powerful series of experiments, Yamaguchi et al. [8] have investigated the behavior of organotypic SCN slices made from neonatal transgenic mice expressing a luciferase reporter driven by an oscillating promoter from the clock gene period-homolog 1 (mPer1); mPer1-luc bioluminescent rhythms in individual cells are measured in long-term culture with a sensitive charge-coupled device (CCD) camera. This technical tour-de-force opens new windows on the workings of SCN tissue at the cellular level, enabling researchers to resolve some previously intractable issues. Among these is the analysis of mutant clock phenotypes. The behavioral arrhythmicity described previously in mutant mice with ‘knocked out’ clock genes could have been due to the loss of individual SCN cellular rhythms – an intracellular defect – or to persistent but desynchronized rhythms among individual cells – a deficit in intercellular coupling. Using their new approach, Yamaguchi et al. have determined that arrhythmicity in www.sciencedirect.com
electrical activity [9] and cyclic expression of mPer1 and mPer2 mRNA [10] of SCN tissue in mice with mutations in two cryptochrome genes (mCry1K/K, mCry2K/K) is in fact due to arrhythmicity of the component cells. SCN cellular synchronization does not mean that all cells express identical phases Importantly, Yamaguchi et al. also have demonstrated that individual SCN cells in normal mouse brain slices show rather large phase differences in the peaks of their bioluminescent rhythms and that these differences persist over repeated cycles (individual cellular periods are similar and stable). The phase order is not a stochastic property of the network because it is restored after cycloheximide is applied, first to stop and then to reset the cellular oscillations to the same initial phase. Furthermore, intercellular phase differences are not an artifact of organotypic culturing of the slices (which lose w70% of their neurons and flatten to a few cell layers thick) because similar differences have been demonstrated in acutely prepared 300 mm mouse slices assayed by an mPer1 green fluorescent protein (mPer1-GFP) reporting system [7], or by electrophysiological methods in rat SCN in vivo [11] or in 500 mm thick acute slices [5]. In general, dorsomedial cells appear to phase-lead (but do not appear to drive) ventrolateral ones in Yamaguchi et al.’s preparation, although other workers have described a lateral-tomedial gradient in Per1 expression [7], and no gradient in electrical activity [5], in acute slices. In any case, it has become clear from all of these studies that the duration of high molecular and high electrical activities of individual SCN cells is shorter than the composite activities of the tissue as a whole (which generally lasts for most of the subjective day) (Figure 1). Good news – there are still more questions than answers What is the functional significance of heterogeneous cellular phases? Can their distribution and clustering [7] or order [12] be reconfigured by light? Could plasticity of phase differences encode a photoperiodic signal [5]? Are they organized as part of the temporal programming of SCN outputs [13]? And what mechanism(s) keep the cells out of phase? Yamaguchi et al. have begun to tackle the last of these questions by applying tetrodotoxin (TTX), which blocks voltage-dependent NaC channels and inhibits the generation of action potentials, to their cultures. At least acutely, rhythms of mPer1-luc bioluminescence persist (as does the rhythm of cytosolic Ca2C levels [14]). But Yamaguchi et al. have now shown that over several cycles of chronic TTX application, the individual cellular bioluminescent rhythms begin to desynchronize from one another. The obvious interpretation is that action potentials synchronize the cells; in their absence, individual cellular oscillations gradually drift and their phase relationships become unstable over succeeding cycles. It is important to note, however, that TTX might also affect other forms of intercellular communication in the SCN (e.g. dye coupling [15]) and that long-term TTX might even unmask compensatory
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TRENDS in Neurosciences Vol.27 No.9 September 2004
Activity
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Figure 1. From oscillating cells to rhythmic tissue. (a) Three cellular oscillations with 4 h phase differences. Cellular activity is linearly spread over the subjective day [circadian time (CT) 0–12 h, with CT6 representing the midday]. The duration of enhanced activity at half-maximum is 5 h for single-unit electrical activity [5]. Numbers on the y-axis are normalized for single-unit activity, and curvature is simple and theoretical. (b) Summed activity of the three cells in (a), resulting in a biphasic activity pattern. (c) Summed activity of 15 cells with a phase distribution linearly spread over subjective day. The composite activity shows a smooth pattern with a single peak at CT6. The most advanced and most delayed units from the population are shown in (a). Plasticity of such phase differences within the suprachiasmatic nucleus could encode a photoperiodic signal [5]. A closer phase relationship between oscillating cells would generate a population signal corresponding to that observed for short winter day-lengths, whereas a wider distribution would create a long summer day-length pattern.
responses that could dynamically change intercellular responsiveness [16]. Surprisingly, the amplitude of the mPer1-luc bioluminescence rhythm dramatically drops over several days of continuous TTX application, and Yamaguchi et al. concluded that action potentials are necessary for maintaining intracellular molecular rhythms. Although not explicitly discussed by the authors, their figures seem to show that diminished amplitudes are also associated with apparently distorted waveforms, raising the possibility that the amplitude effect might be due in part to phase interference between desynchronized cells or even to a toxic effect of chronic TTX on the luciferin–luciferase bioluminescence reaction. It would be useful to know whether the mean period expressed by the attenuated network is altered – that is, whether TTX actually stops, slows or shifts the tissue clock. When TTX is infused into the rat SCN for many days in vivo, entrainment and expression of behavioral rhythmicity is lost but the actual circadian oscillatory mechanism appears to be unaffected [i.e. the phase of the restored rhythm after TTX treatment is that predicted by extrapolation of the phase and period www.sciencedirect.com
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of the original (pre-infusion) rhythm] [17]. Of course, the physiology of the SCN in vitro and in vivo is likely to be different; in vivo, the SCN appears to be more stable (induced phase shifts are smaller in vivo than in vitro), and there is the (theoretical) possibility that extra-SCN brain oscillators might keep time even when the SCN pacemaker is inactivated [18]. Lastly, we call attention to a detail in the imaging study of Yamaguchi et al.’s, although its significance is presently uncertain. The authors note that nearly all of their luminescent cells (99.2% of a total of 1177) exhibit circadian rhythmicity; this was not the case for Quintero et al.’s [7] slices acutely prepared from mPer1-GFP transgenic mice. Furthermore, at least in hamster SCN, there seems to be a subpopulation of calbindin-expressing neurons that do not appear to express circadian rhythmicity (of neither hamster Per1 abundance [19] nor electrical activity [20]). Whether these disagreements are due to reporter sensitivity, species differences, slice thickness or reorganization of circuitry in organotypic slices will require further study. Keeping the membrane in mind The circadian system is complex, and intercellular interactions at the tissue level underlie much of this complexity (not only in the mammalian SCN but also in Drosophila [21]). For the generation of overt rhythmicity, clock genes are necessary but not sufficient (e.g. for vasopressin gene transcription in the rat SCN [22]). Events at the cell membrane lie at the intersection of core oscillatory apparatus of the clock with both its input and output pathways, and a network of interconnected neurons can oscillate without requiring every neuron to be endogenously rhythmic. There are already mutant mice in which absence of the VPAC2 receptor for vasoactive intestinal polypeptide, a neurotransmitter in the retinorecipient subdivision of the SCN, results in a dramatic drop in electrical activity and molecular arrhythmicity [23,24]; in Drosophila too, the state of membrane excitability appears to be key for sustaining free-running intracellular oscillations [25]. Membrane events are crucial to translating rudimentary molecular cycles into a reliable and photo-entrainable tissue pacemaker, and unraveling the dynamic interactions between these system elements should be a challenging problem for years to come. Acknowledgements We thank Erik Herzog and Joseph Miller for helpful discussions. W.J.S. is supported by NINDS grant R01 NS 46605. The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of the NINDS.
References 1 Welsh, D.K. et al. (1995) Individual neurons dissociated from rat suprachiasmatic nucleus express independently phased circadian firing rhythms. Neuron 14, 697–706 2 Liu, C. et al. (1997) Cellular construction of a circadian clock: period determination in the suprachiasmatic nuclei. Cell 91, 855–860 3 Herzog, E.D. et al. (1998) Clock controls circadian period in isolated suprachiasmatic nucleus neurons. Nat. Neurosci. 1, 708–713
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4 Low-Zeddies, S.S. and Takahashi, J.S. (2001) Chimera analysis of the Clock mutation in mice shows that complex cellular integration determines circadian behavior. Cell 105, 25–42 5 Schaap, J. et al. (2003) Heterogeneity of rhythmic suprachiasmatic nucleus neurons: implications for circadian waveform and photoperiodic encoding. Proc. Natl. Acad. Sci. U. S. A. 100, 15994–15999 6 Herzog, E.D. et al. (2004) Temporal precision in the mammalian circadian system: a reliable clock from less reliable neurons. J. Biol. Rhythms 19, 35–46 7 Quintero, J.E. et al. (2003) The biological clock nucleus: a multiphasic oscillator network regulated by light. J. Neurosci. 23, 8070–8076 8 Yamaguchi, S. et al. (2003) Synchronization of cellular clocks in the suprachiasmatic nucleus. Science 302, 1408–1412 9 Albus, H. et al. (2002) Cryptochrome-deficient mice lack circadian electrical activity in the suprachiasmatic nuclei. Curr. Biol. 12, 1130–1133 10 Okamura, H. et al. (1999) Photic induction of mPer1 and mPer2 in Cry-deficient mice lacking a biological clock. Science 286, 2531–2534 11 Saeb-Parsy, K. and Dyball, R.E.J. (2003) Defined cell groups in the rat suprachiasmatic nucleus have different day/night rhythms of singleunit activity in vivo. J. Biol. Rhythms 18, 26–42 12 Nagano, M. et al. (2003) An abrupt shift in the day/night cycle causes desynchrony in the mammalian circadian center. J. Neurosci. 23, 6141–6151 13 Kalsbeek, A. and Buijs, R.M. (2002) Output pathways of the mammalian suprachiasmatic nucleus: coding circadian time by transmitter selection and specific targeting. Cell Tissue Res. 309, 109–118 14 Ikeda, M. et al. (2003) Circadian dynamics of cytosolic and nuclear Ca2C in single suprachiasmatic nucleus neurons. Neuron 38, 253–263 15 Colwell, C.S. (2000) Rhythmic coupling among cells in the suprachiasmatic nucleus. J. Neurobiol. 43, 379–388
16 Turrigiano, G.G. et al. (1998) Activity-dependent scaling of quantal amplitude in neocortical neurons. Nature 391, 892–896 17 Schwartz, W.J. et al. (1987) The suprachiasmatic nuclei contain a tetrodotoxin-resistant circadian pacemaker. Proc. Natl. Acad. Sci. U. S. A. 84, 1694–1698 18 Granados-Fuentes, D. et al. (2004) The suprachiasmatic nucleus entrains, but does not sustain, circadian rhythmicity in the olfactory bulb. J. Neurosci. 24, 615–619 19 Hamada, T. et al. (2002) Expression of Period genes: rhythmic and non-rhythmic compartments of the suprachiasmatic nucleus pacemaker. J. Neurosci. 21, 7742–7750 20 Jobst, E.E. and Allen, C.N. (2002) Calbindin neurons in the hamster suprachiasmatic nucleus do not exhibit a circadian variation in spontaneous firing rate. Eur. J. Neurosci. 16, 2469–2474 21 Pang, Y. et al. (2003) Drosophila free-running rhythms require intercellular communication. PLoS Biol. 1, 32–40 22 Arima, H. et al. (2002) Neuronal activity is required for the circadian rhythm of vasopressin gene transcription in the suprachiasmatic nucleus in vitro. Endocrinology 143, 4165–4171 23 Harmar, A.J. et al. (2002) The VPAC2 receptor is essential for circadian function in the mouse suprachiasmatic nuclei. Cell 109, 497–508 24 Cutler, D.J. et al. (2003) The mouse VPAC2 receptor confers suprachiasmatic nuclei cellular rhythmicity and responsiveness to vasoactive intestinal polypeptide in vitro. Eur. J. Neurosci. 17, 197–204 25 Nitabach, M.N. et al. (2002) Electrical silencing of Drosophila pacemaker neurons stops the free-running circadian clock. Cell 109, 485–495 0166-2236/$ - see front matter Q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.tins.2004.06.016
Schizophrenia genetics: dysbindin under the microscope Matthew A. Benson, Roy V. Sillitoe and Derek J. Blake Department of Pharmacology, University of Oxford, Mansfield Road, Oxford OX1 3QT, UK
It is well established that genetic factors strongly contribute to the susceptibility of an individual to schizophrenia. Straub, Kendler and colleagues have published the first of several articles demonstrating a genetic association between schizophrenia and the gene encoding the dystrobrevin-binding protein dysbindin. Although no mutations in the dysbindin gene have been found, the recent identification of a specific risk haplotype in independent samples provides further evidence that dysbindin is a possible schizophrenia susceptibility gene. Schizophrenia is a highly debilitating, often lifelong illness that affects up to 1% of the population. The fact that schizophrenia has a genetic component has long been established [1] but, in common with many other complex disorders, the genes predisposing to schizophrenia have remained elusive. Recently however, a growing list of Corresponding author: Derek J. Blake ([email protected]). Available online 20 July 2004 www.sciencedirect.com
genes is being linked with susceptibility to the disease [2,3]. Several studies have demonstrated that variation in the gene encoding the dystrobrevin-binding protein dysbindin (DTNBP1) is associated with increased susceptibility to schizophrenia. Dysbindin variation and schizophrenia In the case of DTNBP1, Straub et al. [4] undertook systematic linkage-disequilibrium mapping across the previously linked schizophrenia susceptibility region 6p24–21. They discovered that single nucleotide polymorphisms (SNPs) within the 140 kb DTNBP1 gene were strongly associated with schizophrenia. Twelve SNPs were identified within DTNBP1 and tested for association with schizophrenia in a group of 270 Irish high-density pedigrees [4]. Several of the individual SNPs were significantly associated with schizophrenia and remained so when the data were analysed to include only one affected offspring per nuclear family per extended pedigree. Additionally, Straub et al. found a three-SNP haplotype that was highly significant when restricted
TRENDS in Neurosciences Vol.27 No.9 September 2004
neuronal circuitry. Adult-fly-restricted expression of dominantly acting genes that inhibit evoked neurotransmitter release (e.g. UAS-tetanus toxin light chain [24]) or electrically silence neurons (e.g. UAS-Kir2.1, a genetically engineered KC leak channel [24]) will allow us to better understand the neural circuits underlying complex adult behaviors such as courtship, circadian rhythm, and learning and memory. Another powerful use of Gene-Switch and TARGET would be adult-fly-restricted RNA interference. UAS-driven double-stranded RNA hairpin transgenes [25] combined with TARGET or Gene-Switch could allow neurogeneticists to dissect the role of a gene in adult behavior independent of its role in neurodevelopment. The TARGET and Gene-Switch systems represent a significant technological advance for genetic studies of behavior in Drosophila. Prior to their development, memory researchers had to compromise, selecting either spatially restricted, semi-constitutive gene expression using the standard GAL4-UAS system or temporally controllable, spatially indiscriminate gene expression using the heatshock promoter. Thanks to the efforts of the Davis laboratory, these limitations are now merely memories.
References 1 McGuire, S.E. et al. (2003) Spatiotemporal rescue of memory dysfunction in Drosophila. Science 302, 1765–1768 2 Mao, Z. et al. (2004) Pharmacogenetic rescue in time and space of the rutabaga memory impairment by using Gene-Switch. Proc. Natl. Acad. Sci. U. S. A. 101, 198–203 3 Brand, A.H. and Perrimon, N. (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118, 401–415 4 Duffy, J.B. (2002) GAL4 system in Drosophila: a fly geneticist’s Swiss army knife. Genesis 34, 1–15 5 Lee, T. and Luo, L. (1999) Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22, 451–461 6 Burcin, M.M. et al. (1999) Adenovirus-mediated regulable target gene expression in vivo. Proc. Natl. Acad. Sci. U. S. A. 96, 355–360 7 Osterwalder, T. et al. (2001) A conditional tissue-specific transgene expression system using inducible GAL4. Proc. Natl. Acad. Sci. U. S. A. 98, 12596–12601
513
8 Roman, G. et al. (2001) P[Switch], a system for spatial and temporal control of gene expression in Drosophila melanogaster. Proc. Natl. Acad. Sci. U. S. A. 98, 12602–12607 9 Stebbins, M.J. et al. (2001) Tetracycline-inducible systems for Drosophila. Proc. Natl. Acad. Sci. U. S. A. 98, 10775–10780 10 Bello, B. et al. (1998) Spatial and temporal targeting of gene expression in Drosophila by means of a tetracycline-dependent transactivator system. Development 125, 2193–2202 11 Bieschke, E.T. et al. (1998) Doxycycline-induced transgene expression during Drosophila development and aging. Mol. Gen. Genet. 258, 571–579 12 Han, D.D. et al. (2000) Investigating the function of follicular subpopulations during Drosophila oogenesis through hormonedependent enhancer-targeted cell ablation. Development 127, 573–583 13 Levin, L.R. et al. (1992) The Drosophila learning and memory gene rutabaga encodes a Ca2C/calmodulin-responsive adenylyl cyclase. Cell 68, 479–489 14 Kandel, E.R. et al. (1983) Classical conditioning and sensitization share aspects of the same molecular cascade in Aplysia. Cold Spring Harb. Symp. Quant. Biol. 48, 821–830 15 Livingstone, M.S. et al. (1984) Loss of calcium/calmodulin responsiveness in adenylate cyclase of rutabaga, a Drosophila learning mutant. Cell 37, 205–215 16 Han, P.L. et al. (1992) Preferential expression of the Drosophila rutabaga gene in mushroom bodies, neural centers for learning in insects. Neuron 9, 619–627 17 Zars, T. et al. (2000) Localization of a short-term memory in Drosophila. Science 288, 672–675 18 Corfas, G. and Dudai, Y. (1991) Morphology of a sensory neuron in Drosophila is abnormal in memory mutants and changes during aging. Proc. Natl. Acad. Sci. U. S. A. 88, 7252–7256 19 Balling, A. et al. (1987) Are the structural changes in adult Drosophila mushroom bodies memory traces? Studies on biochemical learning mutants. J. Neurogenet. 4, 65–73 20 McCall, K. and Steller, H. (1997) Facing death in the fly: genetic analysis of apoptosis in Drosophila. Trends Genet. 13, 222–226 21 Roman, G. and Davis, R.L. (2002) Conditional expression of UAS-transgenes in the adult eye with a new gene-switch vector system. Genesis 34, 127–131 22 Rensing, L. and Ruoff, P. (2002) Temperature effect on entrainment, phase shifting, and amplitude of circadian clocks and its molecular bases. Chronobiol. Int. 19, 807–864 23 McGuire, S. et al. (2004) Spatiotemporal gene expression targeting with the TARGET and Gene-Switch systems in Drosophila. Sci. STKE 220, 14 24 White, B. et al. (2001) Molecular genetic approaches to the targeted suppression of neuronal activity. Curr. Biol. 11, R1041–R1053 25 Kalidas, S. and Smith, D.P. (2002) Novel genomic cDNA hybrids produce effective RNA interference in adult Drosophila. Neuron 33, 177–184 0166-2236/$ - see front matter Q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.tins.2004.06.013
Real-time imaging reveals spatiotemporal dynamics of cellular circadian clocks William J. Schwartz1 and Johanna H. Meijer2 1 2
Department of Neurology, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, MA 01655, USA Department of Neurophysiology, Leiden University Medical Centre, 2300 RC Leiden, The Netherlands
Circadian clocks have been localized to discrete sites within the nervous systems of several organisms, and in mammals to the suprachiasmatic nucleus (SCN) in the anterior hypothalamus. The clock in the SCN is Corresponding author: William J. Schwartz ([email protected]). Available online 10 July 2004 www.sciencedirect.com
composed of multiple autonomous single-cell oscillators, and new studies now allow an unprecedented look at their oscillatory activities over repeated cycles in tissue slices in vitro. Initial data reinforce the importance of intercellular membrane events for constructing a functional and reliable tissue pacemaker.
514
Update
TRENDS in Neurosciences Vol.27 No.9 September 2004
To adapt to the once-daily rotation of our planet about its axis, most organisms have evolved to incorporate the geophysical day–night cycle as a genetically determined temporal program. This endogenous, temperaturecompensated timekeeping mechanism – the ‘circadian clock’ – recognizes local time, measures its passage, and contributes to the regulation of homeostasis, seasonal behavior and navigation. Our progress in understanding its neurobiology has been almost unimaginable. About 30 years ago, the suprachiasmatic nucleus (SCN) was first implicated as the site of a master clock, but it has been less than ten years since SCN neurons were discovered to be autonomous cellular circadian oscillators [1]: when cultured at low density, they individually express firing rate rhythms with different circadian periods. Analyses of induced and spontaneous mutations, gene sequence homologies and protein–protein interactions have identified candidate regulatory molecules and biochemical processes that are likely to constitute the basic intracellular oscillatory mechanism. It is believed that genes at the core of the clock function within autoregulatory feedback loops, with nuclear proteins rhythmically suppressing the transcription of their own mRNA. Still mysterious is how individual SCN cells are assembled to create an integrated tissue pacemaker that governs circadian behaviors of whole animals. The genotype-specific circadian period characteristically expressed by animals appears to be an intermediate period that arises from the coupling of multiple SCN cellular oscillators [2–4]; this contrasts with many other rhythmic tissues (e.g. the heart), in which the fastest cells set the rate. Elucidating SCN intercellular interactions will be crucial to understanding circadian properties at the ‘tissue’ level of organization; several attributes of the clock are likely to emerge at this level, such as its regulation of complex behavioral patterns, adaptation to photoperiod [5] and remarkable precision [6]. Time for new technologies Recently there have been exciting advances in developing methods that make possible real-time simultaneous measurements of oscillatory activity over repeated cycles from multiple individual cells in tissue slices [7,8]. In a remarkable and powerful series of experiments, Yamaguchi et al. [8] have investigated the behavior of organotypic SCN slices made from neonatal transgenic mice expressing a luciferase reporter driven by an oscillating promoter from the clock gene period-homolog 1 (mPer1); mPer1-luc bioluminescent rhythms in individual cells are measured in long-term culture with a sensitive charge-coupled device (CCD) camera. This technical tour-de-force opens new windows on the workings of SCN tissue at the cellular level, enabling researchers to resolve some previously intractable issues. Among these is the analysis of mutant clock phenotypes. The behavioral arrhythmicity described previously in mutant mice with ‘knocked out’ clock genes could have been due to the loss of individual SCN cellular rhythms – an intracellular defect – or to persistent but desynchronized rhythms among individual cells – a deficit in intercellular coupling. Using their new approach, Yamaguchi et al. have determined that arrhythmicity in www.sciencedirect.com
electrical activity [9] and cyclic expression of mPer1 and mPer2 mRNA [10] of SCN tissue in mice with mutations in two cryptochrome genes (mCry1K/K, mCry2K/K) is in fact due to arrhythmicity of the component cells. SCN cellular synchronization does not mean that all cells express identical phases Importantly, Yamaguchi et al. also have demonstrated that individual SCN cells in normal mouse brain slices show rather large phase differences in the peaks of their bioluminescent rhythms and that these differences persist over repeated cycles (individual cellular periods are similar and stable). The phase order is not a stochastic property of the network because it is restored after cycloheximide is applied, first to stop and then to reset the cellular oscillations to the same initial phase. Furthermore, intercellular phase differences are not an artifact of organotypic culturing of the slices (which lose w70% of their neurons and flatten to a few cell layers thick) because similar differences have been demonstrated in acutely prepared 300 mm mouse slices assayed by an mPer1 green fluorescent protein (mPer1-GFP) reporting system [7], or by electrophysiological methods in rat SCN in vivo [11] or in 500 mm thick acute slices [5]. In general, dorsomedial cells appear to phase-lead (but do not appear to drive) ventrolateral ones in Yamaguchi et al.’s preparation, although other workers have described a lateral-tomedial gradient in Per1 expression [7], and no gradient in electrical activity [5], in acute slices. In any case, it has become clear from all of these studies that the duration of high molecular and high electrical activities of individual SCN cells is shorter than the composite activities of the tissue as a whole (which generally lasts for most of the subjective day) (Figure 1). Good news – there are still more questions than answers What is the functional significance of heterogeneous cellular phases? Can their distribution and clustering [7] or order [12] be reconfigured by light? Could plasticity of phase differences encode a photoperiodic signal [5]? Are they organized as part of the temporal programming of SCN outputs [13]? And what mechanism(s) keep the cells out of phase? Yamaguchi et al. have begun to tackle the last of these questions by applying tetrodotoxin (TTX), which blocks voltage-dependent NaC channels and inhibits the generation of action potentials, to their cultures. At least acutely, rhythms of mPer1-luc bioluminescence persist (as does the rhythm of cytosolic Ca2C levels [14]). But Yamaguchi et al. have now shown that over several cycles of chronic TTX application, the individual cellular bioluminescent rhythms begin to desynchronize from one another. The obvious interpretation is that action potentials synchronize the cells; in their absence, individual cellular oscillations gradually drift and their phase relationships become unstable over succeeding cycles. It is important to note, however, that TTX might also affect other forms of intercellular communication in the SCN (e.g. dye coupling [15]) and that long-term TTX might even unmask compensatory
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Figure 1. From oscillating cells to rhythmic tissue. (a) Three cellular oscillations with 4 h phase differences. Cellular activity is linearly spread over the subjective day [circadian time (CT) 0–12 h, with CT6 representing the midday]. The duration of enhanced activity at half-maximum is 5 h for single-unit electrical activity [5]. Numbers on the y-axis are normalized for single-unit activity, and curvature is simple and theoretical. (b) Summed activity of the three cells in (a), resulting in a biphasic activity pattern. (c) Summed activity of 15 cells with a phase distribution linearly spread over subjective day. The composite activity shows a smooth pattern with a single peak at CT6. The most advanced and most delayed units from the population are shown in (a). Plasticity of such phase differences within the suprachiasmatic nucleus could encode a photoperiodic signal [5]. A closer phase relationship between oscillating cells would generate a population signal corresponding to that observed for short winter day-lengths, whereas a wider distribution would create a long summer day-length pattern.
responses that could dynamically change intercellular responsiveness [16]. Surprisingly, the amplitude of the mPer1-luc bioluminescence rhythm dramatically drops over several days of continuous TTX application, and Yamaguchi et al. concluded that action potentials are necessary for maintaining intracellular molecular rhythms. Although not explicitly discussed by the authors, their figures seem to show that diminished amplitudes are also associated with apparently distorted waveforms, raising the possibility that the amplitude effect might be due in part to phase interference between desynchronized cells or even to a toxic effect of chronic TTX on the luciferin–luciferase bioluminescence reaction. It would be useful to know whether the mean period expressed by the attenuated network is altered – that is, whether TTX actually stops, slows or shifts the tissue clock. When TTX is infused into the rat SCN for many days in vivo, entrainment and expression of behavioral rhythmicity is lost but the actual circadian oscillatory mechanism appears to be unaffected [i.e. the phase of the restored rhythm after TTX treatment is that predicted by extrapolation of the phase and period www.sciencedirect.com
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of the original (pre-infusion) rhythm] [17]. Of course, the physiology of the SCN in vitro and in vivo is likely to be different; in vivo, the SCN appears to be more stable (induced phase shifts are smaller in vivo than in vitro), and there is the (theoretical) possibility that extra-SCN brain oscillators might keep time even when the SCN pacemaker is inactivated [18]. Lastly, we call attention to a detail in the imaging study of Yamaguchi et al.’s, although its significance is presently uncertain. The authors note that nearly all of their luminescent cells (99.2% of a total of 1177) exhibit circadian rhythmicity; this was not the case for Quintero et al.’s [7] slices acutely prepared from mPer1-GFP transgenic mice. Furthermore, at least in hamster SCN, there seems to be a subpopulation of calbindin-expressing neurons that do not appear to express circadian rhythmicity (of neither hamster Per1 abundance [19] nor electrical activity [20]). Whether these disagreements are due to reporter sensitivity, species differences, slice thickness or reorganization of circuitry in organotypic slices will require further study. Keeping the membrane in mind The circadian system is complex, and intercellular interactions at the tissue level underlie much of this complexity (not only in the mammalian SCN but also in Drosophila [21]). For the generation of overt rhythmicity, clock genes are necessary but not sufficient (e.g. for vasopressin gene transcription in the rat SCN [22]). Events at the cell membrane lie at the intersection of core oscillatory apparatus of the clock with both its input and output pathways, and a network of interconnected neurons can oscillate without requiring every neuron to be endogenously rhythmic. There are already mutant mice in which absence of the VPAC2 receptor for vasoactive intestinal polypeptide, a neurotransmitter in the retinorecipient subdivision of the SCN, results in a dramatic drop in electrical activity and molecular arrhythmicity [23,24]; in Drosophila too, the state of membrane excitability appears to be key for sustaining free-running intracellular oscillations [25]. Membrane events are crucial to translating rudimentary molecular cycles into a reliable and photo-entrainable tissue pacemaker, and unraveling the dynamic interactions between these system elements should be a challenging problem for years to come. Acknowledgements We thank Erik Herzog and Joseph Miller for helpful discussions. W.J.S. is supported by NINDS grant R01 NS 46605. The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of the NINDS.
References 1 Welsh, D.K. et al. (1995) Individual neurons dissociated from rat suprachiasmatic nucleus express independently phased circadian firing rhythms. Neuron 14, 697–706 2 Liu, C. et al. (1997) Cellular construction of a circadian clock: period determination in the suprachiasmatic nuclei. Cell 91, 855–860 3 Herzog, E.D. et al. (1998) Clock controls circadian period in isolated suprachiasmatic nucleus neurons. Nat. Neurosci. 1, 708–713
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4 Low-Zeddies, S.S. and Takahashi, J.S. (2001) Chimera analysis of the Clock mutation in mice shows that complex cellular integration determines circadian behavior. Cell 105, 25–42 5 Schaap, J. et al. (2003) Heterogeneity of rhythmic suprachiasmatic nucleus neurons: implications for circadian waveform and photoperiodic encoding. Proc. Natl. Acad. Sci. U. S. A. 100, 15994–15999 6 Herzog, E.D. et al. (2004) Temporal precision in the mammalian circadian system: a reliable clock from less reliable neurons. J. Biol. Rhythms 19, 35–46 7 Quintero, J.E. et al. (2003) The biological clock nucleus: a multiphasic oscillator network regulated by light. J. Neurosci. 23, 8070–8076 8 Yamaguchi, S. et al. (2003) Synchronization of cellular clocks in the suprachiasmatic nucleus. Science 302, 1408–1412 9 Albus, H. et al. (2002) Cryptochrome-deficient mice lack circadian electrical activity in the suprachiasmatic nuclei. Curr. Biol. 12, 1130–1133 10 Okamura, H. et al. (1999) Photic induction of mPer1 and mPer2 in Cry-deficient mice lacking a biological clock. Science 286, 2531–2534 11 Saeb-Parsy, K. and Dyball, R.E.J. (2003) Defined cell groups in the rat suprachiasmatic nucleus have different day/night rhythms of singleunit activity in vivo. J. Biol. Rhythms 18, 26–42 12 Nagano, M. et al. (2003) An abrupt shift in the day/night cycle causes desynchrony in the mammalian circadian center. J. Neurosci. 23, 6141–6151 13 Kalsbeek, A. and Buijs, R.M. (2002) Output pathways of the mammalian suprachiasmatic nucleus: coding circadian time by transmitter selection and specific targeting. Cell Tissue Res. 309, 109–118 14 Ikeda, M. et al. (2003) Circadian dynamics of cytosolic and nuclear Ca2C in single suprachiasmatic nucleus neurons. Neuron 38, 253–263 15 Colwell, C.S. (2000) Rhythmic coupling among cells in the suprachiasmatic nucleus. J. Neurobiol. 43, 379–388
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Schizophrenia genetics: dysbindin under the microscope Matthew A. Benson, Roy V. Sillitoe and Derek J. Blake Department of Pharmacology, University of Oxford, Mansfield Road, Oxford OX1 3QT, UK
It is well established that genetic factors strongly contribute to the susceptibility of an individual to schizophrenia. Straub, Kendler and colleagues have published the first of several articles demonstrating a genetic association between schizophrenia and the gene encoding the dystrobrevin-binding protein dysbindin. Although no mutations in the dysbindin gene have been found, the recent identification of a specific risk haplotype in independent samples provides further evidence that dysbindin is a possible schizophrenia susceptibility gene. Schizophrenia is a highly debilitating, often lifelong illness that affects up to 1% of the population. The fact that schizophrenia has a genetic component has long been established [1] but, in common with many other complex disorders, the genes predisposing to schizophrenia have remained elusive. Recently however, a growing list of Corresponding author: Derek J. Blake ([email protected]). Available online 20 July 2004 www.sciencedirect.com
genes is being linked with susceptibility to the disease [2,3]. Several studies have demonstrated that variation in the gene encoding the dystrobrevin-binding protein dysbindin (DTNBP1) is associated with increased susceptibility to schizophrenia. Dysbindin variation and schizophrenia In the case of DTNBP1, Straub et al. [4] undertook systematic linkage-disequilibrium mapping across the previously linked schizophrenia susceptibility region 6p24–21. They discovered that single nucleotide polymorphisms (SNPs) within the 140 kb DTNBP1 gene were strongly associated with schizophrenia. Twelve SNPs were identified within DTNBP1 and tested for association with schizophrenia in a group of 270 Irish high-density pedigrees [4]. Several of the individual SNPs were significantly associated with schizophrenia and remained so when the data were analysed to include only one affected offspring per nuclear family per extended pedigree. Additionally, Straub et al. found a three-SNP haplotype that was highly significant when restricted