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Dinis José Silva Afonso Regulation of sleep and circadian rhythms by TARANIS

Universidade do Minho Escola de Ciências da Saúde

Dinis José Silva Afonso

Regulation of sleep and circadian rhythms by TARANIS

UMinho|2016

Governo da República Portuguesa

janeiro de 2016

Universidade do Minho Escola de Ciências da Saúde

Dinis José Silva Afonso Regulation of sleep and circadian rhythms by TARANIS

Tese de Doutoramento em Medicina

Trabalho efectuado sobre a orientação da Professora Doutora Kyunghee Koh e da Professora Doutora Joana Palha

janeiro de 2016

STATEMENT OF INTEGRITY

I hereby declare having conducted my thesis with integrity. I confirm that I have not used plagiarism or any form of falsification of results in the process of the thesis elaboration. I further declare that I have fully acknowledged the Code of Ethical Conduct of the University of Minho.

University of Minho, 04 __/__/____ 01 2016

Assinatura: ________________________________________

“If every single man and woman, child and baby, acts and conducts itself in a known pattern and breaks no walls and differs with no one and experiments in no way and is not sick and does not endanger the ease and peace of mind or steady unbroken flow of the town, then that unit can disappear and never be heard of.”

In “The Pearl” of John Steinbeck

Acknowledgments There are so many people I have to thank. I apologize to those that I do not mention. Otherwise the list would go on forever. I thank my family for their love, support and patience all along my life. I am deeply grateful to my mentor for sharing her knowledge with me, for patiently helping me enter the field of genetics of Drosophila neurobiology, for always clarifying the right path to go, and for not letting me get lost in too many side projects. Thanks, Kyunghee! I thank James Jepson and Angelique Lamaze for helping me understand fly genetics and behavior from the very beginning. I am also very grateful to Daniel Machado, Die Liu, Huihui Pan, Kaitlyn Kallas, Andrea Nam, Kevin Mcvoy, and Alexandra Kenny for all their help. I thank my scientific committee Alexander Mazo, Mark Fortinni and Jayme Jaynes for their thoughtful comments and advice. I thank Joana Palha for transforming a vision into a real project, without which this work wouldn’t have been possible. I thank all students in the MD/PhD program for sharing their own experiences. I also thank the Drosophila community for openly sharing resources. I thank all at the University of Minho and Thomas Jefferson University that made this path into science possible and exciting.

And, I thank the Fundação para a Ciência e Tecnologia for awarding me with the predoctoral fellowship (SFRH/BD/51726/2011).

Resumo

O sono é essencial à vida, e quando perturbado tem um efeito extremamente

prejudicial

na

saúde

humana.

Apesar

do

crescente

reconhecimento da importância do sono, não há muito para oferecer aos pacientes que sofrem de distúrbios do sono. Os tratamentos atuais não produzem os mesmos resultados que o sono normal satisfaz, são altamente inespecíficos e rapidamente tolerados pelos doentes. Encontrar mecanismos reguladores do sono, certamente melhorará os padrões de qualidade de vida. Neste trabalho em Drosophila, identificámos TARANIS (TARA), um gene em homólogo da família de coreguladores de transcrição Trip-Br (SERTAD), necessários para os padrões normais de sono. Através de um rastreio genético, isolámos mutantes de tara com uma redução marcada na quantidade de sono. tara codifica uma proteína do ciclo celular que contém um domínio conservado de ligação à Ciclina-A (CycA). TARA regula os níveis da proteína CycA e interage genetica e fisicamente com a CycA para promover o sono. Além disso, a diminuição dos níveis da proteína Cinase dependente da Ciclina 1 (Cdk1), um parceiro da CycA, resgata o fenótipo de sono nos mutantes de tara e CycA, enquanto o aumento da actividade de Cdk1 reproduziu os fenótipos de tara e CycA, o que sugere que a proteína Cdk1 medeia o papel de TARA e CycA na regulação do sono. Além do papel na regulação do sono, os mutantes tara apresentam um comportamento locomotor arrítmico em condições de escuridão constante (DD). Apesar dos mecanismos pelos quais a TARA regula o comportamento circadiano serem desconhecidos, mostrámos que TARA regula a velocidade do

oscilador molecular circadiano e o débito neuropeptidico dos neurónios pacemaker. À semelhança dos mutantes Clock (Clk), os mutantes tara exibem um padrão de atividade noturna em condições dia/noite (LD). Além disso, a abolição da expressão de tara ou a sua sobreexpressão nos neurónios sLNvs alteram a morfologia da sua projeção dorsal. Em conjunto, estes resultados sugerem que a TARA é importante em vários passos que comunicam as oscilações moleculares para o comportamento locomotor circadiano. No presente trabalho, descrevemos uma nova via genética que controla o sono em Drosophila. Esta via genética é conservada em mamíferos e tem o potencial de regular o sono em seres humanos, por um mecanismo semelhante. Além disso, neste trabalho desvendámos outros fenótipos sob o controlo da TARA, que podem ser usados para projectar experiências futuras direccionadas à compreensão da morfologia neuronal e dos padrões diários de vigília/sono.

Abstract

Sleep is essential for life, and when disturbed has a deleterious effect in human health. Despite the increased awareness of the importance of sleep, not much can be offered to patients suffering from sleep disorders. The current treatments do not produce the same results as real sleep does, are highly unspecific, and quickly tolerated by patients. Finding novel sleep regulatory mechanisms will certainly improve the quality of life. In this work, we identify TARANIS (TARA), a Drosophila homolog of the Trip-Br (SERTAD) family of transcriptional coregulators, as a molecule that is required for normal sleep patterns. Through a forward-genetic screen, we isolated tara as a novel sleep gene associated with a marked reduction in sleep amount. tara encodes a conserved cell-cycle protein that contains a Cyclin A (CycA)-binding homology domain. TARA regulates CycA protein levels and genetically and physically interacts with CycA to promote sleep. Furthermore, decreased levels of Cyclin dependent kinase 1 (Cdk1), a kinase partner of CycA, rescue the short-sleeping phenotype of tara and CycA mutants, while increased Cdk1 activity mimics the tara and CycA phenotypes, suggesting that Cdk1 mediates the role of TARA and CycA in sleep regulation. In addition to the role in the regulation of sleep, tara mutants exhibit arrhythmic locomotor behavior in constant darkness (DD). However, the mechanisms through which TARA regulates circadian behavior are unknown. Here, we show that TARA regulates the speed of the molecular oscillator and the neuropeptide output of the pacemaker neurons. Similarly to Clock (Clk) mutants, tara mutants display a nocturnal pattern of activity in light:dark

conditions (LD). Furthermore, tara knockdown or overexpression in sLNvs alters the morphology of their dorsal projection. Taken together, our data suggest that tara functions in multiple steps that link molecular cycling to overt circadian locomotor behavior. In these work, we describe a novel genetic pathway that controls sleep in Drosophila. This genetic pathway may be conserved in mammals and holds the potential to regulate sleep in humans as well. Furthermore, in this work we unravel other phenotypes under the control of tara, which can be used to design future experiments aimed at understanding neuronal morphology and the daily patterns of wakefulness/sleep.

Contents Chapter 1. Introduction

1

I.

Importance of sleep in health and disease

3

II.

Sleep research across species

6

III.

Circadian regulation of sleep-wake rhythms

10

IV.

taranis, cell cycle genes, and sleep regulation

13

V.

Working hypothesis and Aims

15

Chapter 2. Experimental work

29

Chapter 2.1. TARANIS functions with Cyclin A and Cdk1 in a

31

novel arousal center to control sleep in Drosophila Chapter 2.2 taranis regulates circadian rhythms, nocturnal

59

activity, and neuronal morphology Chapter 3. Discussion, Conclusions and Future Perspectives

95

I.

tara is a novel regulator of sleep

97

II.

tara, CycA, and Cdk1 form a new sleep regulatory signaling

98

pathway III.

A novel arousal circuit modulated by tara, CycA, and Cdk1

101

IV.

tara may control sleep by regulating synaptic structure

103

V.

tara negatively regulates the intrinsic speed of the molecular

104

clock VI.

tara regulates PDF levels

VII.

tara regulates nocturnal activity and neuronal

106 morphology

108

Abbreviations GABA – γ-aminobutyric acid REM – Rapid eye movement NREM – Non-rapid eye movement EEG – Electroencephalography DAM – Drosophila Monitoring System per – period Pdf – Pigment dispersing Factor Gal4 – Transcription factor derived from yeast UAS – Upstream Activating Sequence. GAL4 binds UAS to initiate transcription of downstream sequences LexA – Transcription factor derived from bacteria LexAop – LexA operator. LexA binds LexAop sequence CycA – Cyclin-A cAMP – Cyclic adenosine monophosphate CLK – CLOCK CYC – CYCLE timeless – tim sLNv – small ventral lateral neuron lLNv – large ventral lateral neuron LNd – dorsal lateral neuron DN – dorsal neuron clusters (DN1, DN2, and DN3) LPN – lateral posterior neuron DD – constant darkness conditions LD – light and dark conditions

LL – constant Light conditions CREB – cAMP response element binding protein CRY – CRYPTHOCHROME tara – taranis PHD – Plant homeo domain TRIP-Br – Transcriptional Regulators Interacting with PHD zinc fingers and/or Bromodomains Cdks – Cyclin-dependent kinases PL – pars lateralis MB – Mushroom bodies PI – pars intercerebralis NaChBac – Bacterial Sodium Channel TrpA1 – Warm activated cation channel Cdk1-AF – Cdk1 mutant with elevated kinase activity due to mutations in inhibitory phosphorylation sites TNT-Imp – Inactive form of tetanus neurotoxin TNT-G – Active form of tetanus neurotoxin ZT – Zeitgeber time CT – Circadian time Ub-tara – Ubiquitous promoter upstream of tara DATfmn – Dopamine transporter mutant PP2A – Protein Phosphatase 2A

Chapter 1 ___________________________________________________________________ Introduction

I.

Importance of sleep in health and disease

The brain is complex and generates a wide range of behaviors. Of all the behaviors required for survival, sleep is the most time-consuming, and its loss has many deleterious effects on human physiology [1, 2]. Sleep deprivation disrupts metabolism, increasing glucose intolerance and insulin resistance [3], and shift workers have higher rates of cancer [4]. Sleep deprivation also decreases intellectual ability and motor function, greatly increasing the risk of fatal accidents for sleep deprived drivers. It is estimated that 16.5% of all fatal motor vehicle crashes in the United States are due to drowsy drivers [5]. Finally, sleep disturbances are also highly prevalent in patients with psychiatric diseases, including anxiety, depression, obsessive compulsive disorder and intellectual disability [6, 7], and are an early sign of neurodegenerative diseases in Parkinson’s [8] and Alzheimer’s [2] diseases. Importantly, sleep loss is not simply a symptom of these diseases but a major risk factor in their cause and/or perpetuation [2, 6, 9]. Despite being important for almost all aspects of life, the reasons why we sleep and how sleep is regulated remain largely unknown [10, 11]. While the functions of sleep are not yet clear, several theories have been proposed [12]. First, increasing evidence suggests that sleep is important for memory consolidation and synaptic plasticity [12]. In humans, learning a task leads to an increase in synaptic strength (cortical evoked field potentials) that is followed by a higher slow-wave activity (marker of sleep need) in the subsequent sleep period [12]. Evidence from mice also shows a net increase in synaptic spines during wake and a net decline during sleep [13]. These data are consistent with a theory of sleep function called the synaptic homeostasis hypothesis, which postulates that, during

3

sleep, synapses are downscaled to renormalize neural connectivity [12]. Second, lower energy consumption during sleep is also viewed as a function of sleep [12]. The brain is the most energy-demanding organ in the body, requiring 25% of consumed glucose [12]; however, during slow-wave sleep the energetic demands of the brain are reduced by 25-44% [14]. Finally, a recently proposed function for sleep is brain detoxification where, during sleep, the brain removes waste products to improve cellular homeostasis. Work in rats has demonstrated that during sleep the interstitial space increases by 60%, which leads to a higher exchange of cerebrospinal fluid, improving the removal of potential neurotoxic byproducts that accumulated during wakefulness [15]. Given the evidence supporting these various hypotheses, it is possible that sleep serves multiple functions. That sleep is essential for life and that its disorders must be treated is evident. However, the currently available treatments for sleep are highly unsatisfactory. The widely used drugs in the treatment of insomnia, benzodiazepines, are unspecific and have a high risk of dependence [16]. Benzodiazepines bind to GABAA receptors and potentiate the postsynaptic response to γ-aminobutyric acid (GABA), decreasing neuronal activity globally. However, reducing neuronal activity is not the same as inducing sleep, as the brain electrical activity during rapid eye movement (REM) sleep is as high as during wakefulness [17]. In addition, under the influence of the benzodiazepines, the overall sleep architecture is altered, with less REM sleep and less N3 non-rapid eye movement sleep (NREM) sleep (slow-wave sleep). The current limitations of the available treatments for sleep disorders highlight the need for the discovery of new sleep regulatory mechanisms that can be used as novel drug targets. In addition, determining the molecular mechanisms of sleep may

4

contribute to our understanding of related processes including memory and cognition.

5

II.

Sleep research across species

Sleep is conserved from flies to humans [18] and Drosophila shares with humans several important features of sleep [19, 20]. During sleep, flies become immobile and adopt a specific sleep posture. Similar to humans, flies sleep mostly at night and exhibit a compensatory response to sleep deprivation. In addition, drugs that promote wakefulness in humans such as caffeine [19], modafinil [21], and amphetamine [22], also promote arousal in flies, suggesting that the underlying regulatory mechanisms may be conserved from flies to humans. In humans, sleep is defined on the basis of brain electrical activity as detected by electrodes placed on the scalp. Polysomnographic profiles identify two states of sleep: REM and NREM sleep. NREM sleep is further subdivided into three stages, N1 to N3, characterized by increasing arousal thresholds and slowing of cortical electroencephalographic (EEG) activity. Sleep in Drosophila is typically monitored using the Drosophila Monitoring System (DAM) or video recording, which allows automated quantification of locomotor activity [23, 24]. After 5 min of immobility, flies exhibit increased arousal thresholds, which provides the rationale for the commonly accepted definition of sleep as immobility lasting 5 min or longer [19]. In contrast to human research, which is expensive with significant technical and ethical limitations, Drosophila enables the use of sophisticated genetic tools that, combined with a fast life cycle and easy handling, allow systematic and unbiased screens for new genes, neurons, and circuits. Mutational analysis and targeted manipulations of neuronal excitability have the potential to uncover novel signaling pathways and regulatory mechanisms that underlie a specific process or behavior. Furthermore, Drosophila has the advantage of less genomic redundancy and

6

compensation [25]. A simpler genome is useful because a single genetic alteration is more likely to produce a phenotype that can be observed and quantified; it is a much easier entry point for new signaling pathways. Importantly, Drosophila has functional orthologs of ~75% of the human disease-related genes [26]. Therefore it is not surprising Drosophila is gaining momentum in the study of various human diseases, ranging from intellectual disability [7], depressive syndromes and bipolar disorders [27]

to

cardiomyopathies

[28],

and

neurodegenerative

diseases,

including

Parkinson’s, Alzheimer’s, and amyotrophic lateral sclerosis [29]. The genetic tools provided by the fruit fly Drosophila melanogaster allow the dissection of complex and largely unknown behaviors such as sleep. Forward genetic screens allow unbiased approaches to be directed at the discovery of new regulatory mechanisms and signaling pathways underlying complex processes. In forward genetic screens, random, genome-wide mutations are systematically screened in search of a phenotypic alteration of interest. Perhaps one of the most significant contributions to behavioral neuroscience was the discovery of period (per) mutants in 1971 through a forward genetic screen [30]. This discovery was remarkable because it caused a dramatic change in our view of how genes control behaviors. Until then scientists were resistant to accept that a single gene would by itself control a behavior, claiming “genetic architecture of behavior is complex and multigenic” [31], and were highly skeptical of fly studies [32, 33]. The discovery of per mutants and the work that followed led to the discovery of the molecular clock, highlighting the power of the genetic tools available in Drosophila. In a little over 100 years, research in Drosophila has contributed tremendously to the understanding of vertebrate neuroscience [34]. The fly community has developed a wide variety of tools ranging from libraries of Drosophila mutants with

7

genome-wide random transposon insertions to binary gene expression systems including Gal4/UAS [35], derived from yeast, and LexA/LexAop [36, 37], derived from bacteria. Binary systems are composed of a transcription factor and a DNA sequence to which the transcription factor binds, leading to the expression of whatever sequence is downstream. The binary systems are powerful genetic tools that allow targeted alterations of gene expression in living organisms, with spatial and temporal control. The phenotypic consequences of these specific manipulations can be monitored through high-throughput and automated behavioral paradigms. Since the first description of a sleep state in Drosophila in 2000 [19, 20], the field has identified a number of genes that play important roles in sleep. The functions of the diverse array of genes include: neurotransmission (dopamine, octopamine, serotonin, and GABA pathway genes) [38-42], neuropeptide signaling (diuretic hormone 31, Pigment-dispersing factor (Pdf), short neuropeptide F precursor, Sex peptide receptor) [43-45], regulation of neuronal excitability (Shaker, quiver, wide awake, and nicotinic Acetylcholine Receptor α4) [46-49], protein degradation (insomniac) [42, 50], control of cell cycle progression (Cyclin-A (CycA) and Regulator of Cyclin-A) [51], intracellular signaling (protein kinase A, cAMP response element binding protein (CREB), crossveinless c) [52-55], synaptic structure (Neurexin 1, Neuroligin 4, Fmr1) [56-58], and chromatin remodeling (Tat interactive protein 60kDa) [59]. Given the variety of genes involved, it is important to understand how these genes interact with each other to regulate sleep and not simply how each by itself affects sleep. The mechanistic insights gained in Drosophila may help elucidate the molecular basis of sleep regulation in vertebrates. Shaker, one of the first genes identified in sleep regulation, mediates a voltage-activated fast-inactivating IA current

8

that has a major role in membrane repolarization in vertebrates [60]. Importantly, mice lacking the closest mammalian homolog of Shaker, the Kv1.2 channel, also sleep less [61]. Another important insight gained in Drosophila that was successfully translated to mammals was the identification of the cyclic adenosine monophosphate (cAMP)-CREB pathway that is important for learning, memory, and intellectual ability. Initial work in Drosophila identified rutabaga (rut), which encodes a Ca2+/calmodulin-activated adenylyl cyclase [62] and dunce (dnc), which encodes a cAMP-specific phosphodiesterase [63]. Remarkably, several genes underlying intellectual disability, which is a common neurodevelopmental disorder affecting 3% of the population [64], disrupt the cAMP-CREB pathway [7]. Furthermore, most genes affected by sleep deprivation have a cAMP-responsive element [65], and sleep deprivation impairs cAMP-CREB signaling [66], which may explain the memory deficits observed with sleep loss [67].

9

III.

Circadian regulation of sleep-wake rhythms

Sleep is regulated by two main mechanisms [18]: a circadian mechanism that concentrates sleep to a specific period of the day and a homeostatic mechanism that controls sleep amount. Life on earth has evolved to anticipate recurring changes in light and temperature driven by earth’s spin. Animals have adapted their behavior to these changes, and sleep is one of the most prominent behaviors that exhibit circadian rhythms. The circadian control of behavior is mediated through complex molecular oscillators that can maintain daily cycling even in constant environmental conditions. Interestingly, these molecular mechanisms are highly conserved across evolution, highlighting its selective advantage. At the core of the molecular oscillators are transcriptional-translational feedback loops, where transcriptional activators drive the expression of their own repressors. In Drosophila, the expression of the per and timeless (tim) transcripts are driven by CLOCK (CLK) and CYCLE (CYC), and PER and TIM suppress the transcriptional activity of CLK and CYC. Additional feedback loops, posttranscriptional regulations, and post-translational modifications contribute to the precise time course of molecular oscillations [23]. These molecular oscillations take place in ~150 neurons in the adult Drosophila brain which can be divided into several symmetric clusters of distinct anatomy and physiology: the small and large ventral lateral neurons (sLNv and lLNv, respectively), the dorsal lateral neurons (LNd), three dorsal neuron clusters (DN1, DN2, and DN3) and the lateral posterior neurons (LPN) [23]. The sLNvs are necessary and sufficient to drive circadian locomotor rhythms in constant darkness conditions (DD) [68, 69]. In light and dark conditions (LD), the sLNvs drive the morning peak of activity while the LNds and the 5th sLNv drive the

10

evening peak [68, 69]. The DN1 neurons receive input from the sLNvs and integrate information about light and temperature [70, 71]. The lLNvs communicate with the sLNvs through PDF and are waking promoting cells. The function of the DN2 and DN3 clusters is less well understood. An important feature of the circadian mechanism is its ability to synchronize itself to the external environmental conditions. The circadian mechanism can be divided into three components: the input pathway that relays information from the external environment to the core molecular clock machinery, the molecular clock itself, and the output pathway that translates the molecular clock oscillations to overt behaviors and physiologic functions. The input pathway and the core molecular machinery are fairly well dissected [72, 73]. Briefly, the entrainment of the Drosophila clock to light happens through the light-induced degradation of TIM [74-76], that is transmitted to TIM through the blue-light photoreceptor CRYPTHOCHROME (CRY) [77]. TIM and CRY are degraded by the proteasome in response to light, and the E3 ubiquitin ligase, JETLAG, mediates the light-dependent degradation of TIM [78]. Much less is understood about the output pathway and it remains unclear how circadian signals are integrated with other signals to control behavior. In mammals, the molecular timekeeper is remarkably similar to that of Drosophila. Mammalian CLOCK and BMAL1 positively regulate the mammalian period genes (per1, per2, and per3) and the Cryptochrome genes (Cry1 and Cry2). PER and CRY proteins dimerize, form a complex and translocate to the nucleus, repressing the transcriptional activity of CLOCK and BMAL1 [79]. In mammals, the suprachiasmatic nucleus, composed of ~20,000 neurons, acts as a master pacemaker for circadian rhythms.

11

Peripheral clocks have been observed throughout the body of mammals [80], and it is estimated that ~10% of their genes have circadian expression [81, 82]. Therefore, it is not surprising that the circadian mechanism contributes to many aspects of health and disease. Unstable angina, myocardial infarction, and sudden cardiac death have been reported to occur more frequently in the morning within a few hours of awakening [83], and it has been postulated that circadian genes have a major role in synchronizing cardiomyocyte metabolic activity. Asthma symptoms are higher at night and the degree of bronchoconstriction of the airway tree has a circadian profile [84]. Moreover, patients with bipolar disease have altered circadian rhythmicity

and

lithium

therapy

appears

to

be

beneficial

through

the

resynchronization of circadian rhythms [85]. It is likely that Drosophila studies will continue to enlighten our understanding of how sleep and circadian rhythms impact health and disease.

12

IV.

taranis, cell cycle genes, and sleep regulation

Through a forward genetic screen, we discovered a novel mutant with reduced sleep amount and disrupted rhythmicity in locomotor activity. Genotypic mapping of this new mutant revealed that the locus of the previously identified taranis (tara) gene was affected. Previous work described TARA as a transcriptional co-regulator [86]. tara encodes two isoforms, TARA-A and TARA-B, that are functionally interchangeable; both isoforms have several evolutionarily conserved domains: a CYCLIN-A-binding homology domain, a SERTA motif with unknown function, a plant homeo domain (PHD)-bromo binding domain, and a C-terminal domain implicated in transcriptional regulation [86]. More recently, tara was demonstrated to play a role in neuronal development through its interaction with E2f/Dp1 cell cycle regulators [87]. A recent study has proposed that tara has a role in tissue regeneration by preventing other signaling pathways to alter the fate of the regenerating cells [88], while another recent study showed that tara is upregulated in the Drosophila gut after pathogen-induced wounds [89]. However, no behavioral role has previously been assigned to tara. Mammalian homologs of TARA, the TRIP-Br (Transcriptional Regulators Interacting with PHD zinc fingers and/or Bromodomains) family of proteins, also play important roles in cell cycle control through their interaction with E2f/Dp1 [90] and direct binding of Cyclin-D/Cdk4 [91], which shows tara functions are conserved from flies to humans. Importantly, additional roles have been described for Trip-Br molecules including: regulation of lipolysis by Trip-Br2 [92] and function of pancreatic β-cells byTrip-Br1 [93]. However, it is not known whether the tara mammalian homologs regulate sleep or circadian rhythms.

13

TARA and its mammalian homologs have a CycA-binding homology domain, which is relevant given that CycA and its regulator Rca1 control sleep in Drosophila [51]. These observations suggest that tara and CycA may work in the same molecular pathway to regulate sleep. Cyclins regulate cell cycle progression through its modulation of cyclin-dependent kinases (Cdks). Previous work showed that CycA can physically interact with Cdk1 [94] which also raises the possibility that Cdk1 may have a role in sleep as well. Importantly, it has been shown that cell cycle regulators have additional functions in adult neurons. Cyclin E, for instance, plays a role in memory formation and synaptic plasticity [95]. In addition, Cdk4 and Cyclin-B knockdown in bas1 (bang-sensitive1) mutants rescue duration of seizures [96] suggesting these cell-cycle regulators also modulate ion channel activity in the adult Drosophila brain. In humans with temporal lobe epilepsy, Cyclin-B1 is upregulated in the hypothalamus [97], further demonstrating a role beyond cell cycle control.

14

V.

Working hypothesis and aims

Several genes involved in sleep regulation have been found through genomewide forward genetic screens [46, 47, 51]. tara was found in a forward-genetic screen for short-sleeping mutants. The overall goal of this thesis project was to investigate the role of tara in sleep and in circadian behaviors.

Specific aims:

1. Characterize sleep and circadian rhythms in tara mutants; 2. Identify the anatomical loci in which tara regulates sleep and rhythm behavior; 3. Identify the molecular pathways in which TARA regulates sleep and circadian rhythms.

In here, we describe how tara regulates sleep and circadian rhythms through distinct molecular mechanisms in separate neuronal populations. In chapter 2.1 we describe a novel role for tara in sleep regulation through its interaction with CycA and Cdk1 in a novel arousal center. In chapter 2.2 we describe a role for tara in circadian regulation through its modulatory action over the intrinsic speed of the molecular clock and PDF levels. Chapter 3 presents an overall discussion of the results with future perspectives.

15

VI.

References

1.

Grandner, M.A., Jackson, N.J., Pak, V.M., and Gehrman, P.R. (2012). Sleep disturbance is associated with cardiovascular and metabolic disorders. Journal of Sleep Research 21, 427-433.

2.

Palma, J.-A., Urrestarazu, E., and Iriarte, J. (2013). Sleep loss as risk factor for neurologic disorders: A review. Sleep Medicine 14, 229-236.

3.

Buxton, O.M., Cain, S.W., O'Connor, S.P., Porter, J.H., Duffy, J.F., Wang, W., Czeisler, C.A., and Shea, S.A. (2012). Adverse metabolic consequences in humans of prolonged sleep restriction combined with circadian disruption. Science translational medicine 4, 129ra143.

4.

Wang, X.S., Armstrong, M.E., Cairns, B.J., Key, T.J., and Travis, R.C. (2011). Shift work and chronic disease: the epidemiological evidence. Occupational medicine 61, 78-89.

5.

Tefft, B.C. (2012). Prevalence of motor vehicle crashes involving drowsy drivers, United States, 1999-2008. Accident; analysis and prevention 45, 180186.

6.

Asarnow, L.D., Soehner, A.M., and Harvey, A.G. (2014). Basic sleep and circadian science as building blocks for behavioral interventions: a translational approach for mood disorders. Behavioral neuroscience 128, 360370.

7.

Androschuk, A., Al-Jabri, B., and Bolduc, F. (2015). From learning to memory: what flies can tell us about intellectual disability treatment. Frontiers in Psychiatry 6.

16

8.

Iranzo, A., Molinuevo, J.L., Santamaría, J., Serradell, M., Martí, M.J., Valldeoriola, F., and Tolosa, E. (2006). Rapid-eye-movement sleep behaviour disorder as an early marker for a neurodegenerative disorder: a descriptive study. Lancet Neurology 5, 572-577.

9.

Chung, K.H., Li, C.Y., Kuo, S.Y., Sithole, T., Liu, W.W., and Chung, M.H. (2015). Risk of psychiatric disorders in patients with chronic insomnia and sedative-hypnotic prescription: a nationwide population-based follow-up study. Journal of clinical sleep medicine : JCSM : official publication of the American Academy of Sleep Medicine 11, 543-551.

10.

Frank, M.G. (2006). The mystery of sleep function: current perspectives and future directions. Reviews in the neurosciences 17, 375-392.

11.

Sehgal, A., and Mignot, E. (2011). Genetics of sleep and sleep disorders. Cell 146, 194-207.

12.

Tononi, G., and Cirelli, C. (2014). Sleep and the price of plasticity: from synaptic and cellular homeostasis to memory consolidation and integration. Neuron 81, 12-34.

13.

Maret, S., Faraguna, U., Nelson, A.B., Cirelli, C., and Tononi, G. (2011). Sleep and waking modulate spine turnover in the adolescent mouse cortex. Nature neuroscience 14, 1418-1420.

14.

Madsen, P.L., and Vorstrup, S. (1991). Cerebral blood flow and metabolism during sleep. Cerebrovascular and brain metabolism reviews 3, 281-296.

15.

Xie, L., Kang, H., Xu, Q., Chen, M.J., Liao, Y., Thiyagarajan, M., O'Donnell, J., Christensen, D.J., Nicholson, C., Iliff, J.J., et al. (2013). Sleep drives metabolite clearance from the adult brain. Science 342, 373-377.

17

16.

Janhsen, K., Roser, P., and Hoffmann, K. (2015). The problems of long-term treatment with benzodiazepines and related substances. Deutsches Arzteblatt international 112, 1-7.

17.

Schmidt, M.H. (2014). The energy allocation function of sleep: a unifying theory of sleep, torpor, and continuous wakefulness. Neuroscience and biobehavioral reviews 47, 122-153.

18.

Campbell, S.S., and Tobler, I. (1984). Animal sleep: a review of sleep duration across phylogeny. Neuroscience and biobehavioral reviews 8, 269-300.

19.

Hendricks, J.C., Finn, S.M., Panckeri, K.A., Chavkin, J., Williams, J.A., Sehgal, A., and Pack, A.I. (2000). Rest in Drosophila is a sleep-like state. Neuron 25, 129-138.

20.

Shaw, P.J., Cirelli, C., Greenspan, R.J., and Tononi, G. (2000). Correlates of sleep and waking in Drosophila melanogaster. Science 287, 1834-1837.

21.

Hendricks, J.C., Kirk, D., Panckeri, K., Miller, M.S., and Pack, A.I. (2003). Modafinil maintains waking in the fruit fly drosophila melanogaster. Sleep 26, 139-146.

22.

Andretic, R., van Swinderen, B., and Greenspan, R.J. (2005). Dopaminergic modulation of arousal in Drosophila. Current biology : CB 15, 1165-1175.

23.

Allada, R., and Chung, B.Y. (2010). Circadian organization of behavior and physiology in Drosophila. Annu Rev Physiol 72, 605-624.

24.

Gilestro, G.F. (2012). Video tracking and analysis of sleep in Drosophila melanogaster. Nature protocols 7, 995-1007.

25.

Cirelli, C. (2009). The genetic and molecular regulation of sleep: from fruit flies to humans. Nature reviews. Neuroscience 10, 549-560.

18

26.

Reiter, L.T., Potocki, L., Chien, S., Gribskov, M., and Bier, E. (2001). A systematic analysis of human disease-associated gene sequences in Drosophila melanogaster. Genome research 11, 1114-1125.

27.

Zordan, M.A., and Sandrelli, F. (2015). Circadian clock dysfunction and psychiatric disease: could fruit flies have a say? Frontiers in Neurology 6.

28.

Ocorr, K., Vogler, G., and Bodmer, R. (2014). Methods to assess Drosophila heart development, function and aging. Methods 68, 265-272.

29.

Gama Sosa, M.A., De Gasperi, R., and Elder, G.A. (2012). Modeling human neurodegenerative diseases in transgenic systems. Human genetics 131, 535-563.

30.

Konopka, R.J., and Benzer, S. (1971). Clock mutants of Drosophila melanogaster. Proceedings of the National Academy of Sciences of the United States of America 68, 2112-2116.

31.

Greenspan, R.J. (2008). The origins of behavioral genetics. Current biology : CB 18, R192-198.

32.

Anderson, D., and Brenner, S. (2008). Obituary: Seymour Benzer (19212007). Nature 451, 139.

33.

Jan, Y.N., and Jan, L. (2008). Retrospective: Seymour Benzer (1921-2007). Science 319, 45.

34.

Bellen, H.J., Tong, C., and Tsuda, H. (2010). 100 years of Drosophila research and its impact on vertebrate neuroscience: a history lesson for the future. Nature reviews. Neuroscience 11, 514-522.

35.

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.

19

36.

Lai, S.L., and Lee, T. (2006). Genetic mosaic with dual binary transcriptional systems in Drosophila. Nature neuroscience 9, 703-709.

37.

Yagi, R., Mayer, F., and Basler, K. (2010). Refined LexA transactivators and their use in combination with the Drosophila Gal4 system. Proceedings of the National Academy of Sciences of the United States of America 107, 1616616171.

38.

Kume, K., Kume, S., Park, S.K., Hirsh, J., and Jackson, F.R. (2005). Dopamine is a regulator of arousal in the fruit fly. The Journal of neuroscience : the official journal of the Society for Neuroscience 25, 7377-7384.

39.

Crocker, A., and Sehgal, A. (2008). Octopamine regulates sleep in drosophila through

protein

kinase

A-dependent

mechanisms.

The

Journal

of

neuroscience : the official journal of the Society for Neuroscience 28, 93779385. 40.

Yuan, Q., Joiner, W.J., and Sehgal, A. (2006). A sleep-promoting role for the Drosophila serotonin receptor 1A. Current biology : CB 16, 1051-1062.

41.

Agosto, J., Choi, J.C., Parisky, K.M., Stilwell, G., Rosbash, M., and Griffith, L.C. (2008). Modulation of GABAA receptor desensitization uncouples sleep onset and maintenance in Drosophila. Nature neuroscience 11, 354-359.

42.

Stavropoulos, N., and Young, M.W. (2011). insomniac and Cullin-3 regulate sleep and wakefulness in Drosophila. Neuron 72, 964-976.

43.

Kunst, M., Hughes, M.E., Raccuglia, D., Felix, M., Li, M., Barnett, G., Duah, J., and Nitabach, M.N. (2014). Calcitonin gene-related Peptide neurons mediate sleep-specific circadian output in Drosophila. Current biology : CB 24, 2652-2664.

20

44.

Shang, Y., Donelson, Nathan C., Vecsey, Christopher G., Guo, F., Rosbash, M., and Griffith, Leslie C. (2013). Short Neuropeptide F Is a Sleep-Promoting Inhibitory Modulator. Neuron 80, 171-183.

45.

Parisky, K.M., Agosto, J., Pulver, S.R., Shang, Y., Kuklin, E., Hodge, J.J., Kang, K., Liu, X., Garrity, P.A., Rosbash, M., et al. (2008). PDF cells are a GABA-responsive wake-promoting component of the Drosophila sleep circuit. Neuron 60, 672-682.

46.

Cirelli, C., Bushey, D., Hill, S., Huber, R., Kreber, R., Ganetzky, B., and Tononi, G. (2005). Reduced sleep in Drosophila Shaker mutants. Nature 434, 1087-1092.

47.

Koh, K., Joiner, W.J., Wu, M.N., Yue, Z., Smith, C.J., and Sehgal, A. (2008). Identification of SLEEPLESS, a sleep-promoting factor. Science 321, 372376.

48.

Liu, S., Lamaze, A., Liu, Q., Tabuchi, M., Yang, Y., Fowler, M., Bharadwaj, R., Zhang, J., Bedont, J., Blackshaw, S., et al. (2014). WIDE AWAKE Mediates the Circadian Timing of Sleep Onset. Neuron 82, 151-166.

49.

Shi, M., Yue, Z., Kuryatov, A., Lindstrom, J.M., and Sehgal, A. (2014). Identification of Redeye, a new sleep-regulating protein whose expression is modulated by sleep amount, Volume 3.

50.

Pfeiffenberger, C., and Allada, R. (2012). Cul3 and the BTB adaptor insomniac are key regulators of sleep homeostasis and a dopamine arousal pathway in Drosophila. PLoS Genet 8, e1003003.

51.

Rogulja, D., and Young, M.W. (2012). Control of sleep by cyclin A and its regulator. Science 335, 1617-1621.

21

52.

Donlea, J.M., Pimentel, D., and Miesenbock, G. (2014). Neuronal machinery of sleep homeostasis in Drosophila. Neuron 81, 860-872.

53.

Crocker, A., and Sehgal, A. (2008). Octopamine Regulates Sleep in Drosophila through Protein Kinase A-Dependent Mechanisms. The Journal of Neuroscience 28, 9377-9385.

54.

Joiner, W.J., Crocker, A., White, B.H., and Sehgal, A. (2006). Sleep in Drosophila is regulated by adult mushroom bodies. Nature 441, 757-760.

55.

Hendricks, J.C., Williams, J.A., Panckeri, K., Kirk, D., Tello, M., Yin, J.C., and Sehgal, A. (2001). A non-circadian role for cAMP signaling and CREB activity in Drosophila rest homeostasis. Nature neuroscience 4, 1108-1115.

56.

Larkin, A., Chen, M.Y., Kirszenblat, L., Reinhard, J., van Swinderen, B., and Claudianos, C. (2015). Neurexin-1 regulates sleep and synaptic plasticity in Drosophila melanogaster. The European journal of neuroscience.

57.

Li, Y., Zhou, Z., Zhang, X., Tong, H., Li, P., Zhang, Z.C., Jia, Z., Xie, W., and Han, J. (2013). Drosophila Neuroligin 4 Regulates Sleep through Modulating GABA Transmission. The Journal of Neuroscience 33, 15545-15554.

58.

Bushey, D., Tononi, G., and Cirelli, C. (2009). The Drosophila fragile X mental retardation gene regulates sleep need. The Journal of neuroscience : the official journal of the Society for Neuroscience 29, 1948-1961.

59.

Pirooznia, S.K., Chiu, K., Chan, M.T., Zimmerman, J.E., and Elefant, F. (2012). Epigenetic regulation of axonal growth of Drosophila pacemaker cells by histone acetyltransferase tip60 controls sleep. Genetics 192, 1327-1345.

60.

Schwarz, T.L., Tempel, B.L., Papazian, D.M., Jan, Y.N., and Jan, L.Y. (1988). Multiple potassium-channel components are produced by alternative splicing at the Shaker locus in Drosophila. Nature 331, 137-142.

22

61.

Douglas, C.L., Vyazovskiy, V., Southard, T., Chiu, S.Y., Messing, A., Tononi, G., and Cirelli, C. (2007). Sleep in Kcna2 knockout mice. BMC biology 5, 42.

62.

Levin, L.R., Han, P.L., Hwang, P.M., Feinstein, P.G., Davis, R.L., and Reed, R.R. (1992). The Drosophila learning and memory gene rutabaga encodes a Ca2+/Calmodulin-responsive adenylyl cyclase. Cell 68, 479-489.

63.

Nighorn, A., Healy, M.J., and Davis, R.L. (1991). The cyclic AMP phosphodiesterase encoded by the Drosophila dunce gene is concentrated in the mushroom body neuropil. Neuron 6, 455-467.

64.

Yeargin-Allsopp, M., and Boyle, C. (2002). Overview: the epidemiology of neurodevelopmental

disorders.

Mental

retardation

and

developmental

disabilities research reviews 8, 113-116. 65.

Wang, H., Liu, Y., Briesemann, M., and Yan, J. (2010). Computational analysis of gene regulation in animal sleep deprivation. Physiological genomics 42, 427-436.

66.

Vecsey, C.G., Baillie, G.S., Jaganath, D., Havekes, R., Daniels, A., Wimmer, M., Huang, T., Brown, K.M., Li, X.Y., Descalzi, G., et al. (2009). Sleep deprivation impairs cAMP signalling in the hippocampus. Nature 461, 11221125.

67.

Abel, T., Havekes, R., Saletin, J.M., and Walker, M.P. (2013). Sleep, plasticity and memory from molecules to whole-brain networks. Current biology : CB 23, R774-788.

68.

Grima, B., Chelot, E., Xia, R., and Rouyer, F. (2004). Morning and evening peaks of activity rely on different clock neurons of the Drosophila brain. Nature 431, 869-873.

23

69.

Stoleru, D., Peng, Y., Agosto, J., and Rosbash, M. (2004). Coupled oscillators control morning and evening locomotor behaviour of Drosophila. Nature 431, 862-868.

70.

Zhang, L., Chung, B.Y., Lear, B.C., Kilman, V.L., Liu, Y., Mahesh, G., Meissner, R.A., Hardin, P.E., and Allada, R. (2010). DN1(p) circadian neurons coordinate acute light and PDF inputs to produce robust daily behavior in Drosophila. Current biology : CB 20, 591-599.

71.

Zhang, Y., Liu, Y., Bilodeau-Wentworth, D., Hardin, P.E., and Emery, P. (2010). Light and temperature control the contribution of specific DN1 neurons to Drosophila circadian behavior. Current biology : CB 20, 600-605.

72.

Sehgal, A., Joiner, W., Crocker, A., Koh, K., Sathyanarayanan, S., Fang, Y., Wu, M., Williams, J.A., and Zheng, X. (2007). Molecular analysis of sleep: wake cycles in Drosophila. Cold Spring Harbor symposia on quantitative biology 72, 557-564.

73.

Dubruille, R., and Emery, P. (2008). A Plastic Clock: How Circadian Rhythms Respond to Environmental Cues in Drosophila. Molecular neurobiology 38, 129-145.

74.

Hunter-Ensor, M., Ousley, A., and Sehgal, A. (1996). Regulation of the Drosophila protein timeless suggests a mechanism for resetting the circadian clock by light. Cell 84, 677-685.

75.

Myers, M.P., Wager-Smith, K., Rothenfluh-Hilfiker, A., and Young, M.W. (1996). Light-induced degradation of TIMELESS and entrainment of the Drosophila circadian clock. Science 271, 1736-1740.

76.

Zeng, H., Qian, Z., Myers, M.P., and Rosbash, M. (1996). A light-entrainment mechanism for the Drosophila circadian clock. Nature 380, 129-135.

24

77.

Stanewsky, R., Kaneko, M., Emery, P., Beretta, B., Wager-Smith, K., Kay, S.A., Rosbash, M., and Hall, J.C. (1998). The cryb mutation identifies cryptochrome as a circadian photoreceptor in Drosophila. Cell 95, 681-692.

78.

Koh, K., Zheng, X., and Sehgal, A. (2006). JETLAG resets the Drosophila circadian clock by promoting light-induced degradation of TIMELESS. Science 312, 1809-1812.

79.

Mohawk, J.A., Green, C.B., and Takahashi, J.S. (2012). Central and peripheral circadian clocks in mammals. Annual review of neuroscience 35, 445-462.

80.

Yoo, S.H., Yamazaki, S., Lowrey, P.L., Shimomura, K., Ko, C.H., Buhr, E.D., Siepka,

S.M.,

Hong,

H.K.,

Oh,

W.J.,

Yoo,

O.J.,

et

al.

(2004).

PERIOD2::LUCIFERASE real-time reporting of circadian dynamics reveals persistent circadian oscillations in mouse peripheral tissues. Proceedings of the National Academy of Sciences of the United States of America 101, 53395346. 81.

Doherty, C.J., and Kay, S.A. (2010). Circadian control of global gene expression patterns. Annual review of genetics 44, 419-444.

82.

Panda, S., Antoch, M.P., Miller, B.H., Su, A.I., Schook, A.B., Straume, M., Schultz, P.G., Kay, S.A., Takahashi, J.S., and Hogenesch, J.B. (2002). Coordinated transcription of key pathways in the mouse by the circadian clock. Cell 109, 307-320.

83.

Virag, J.A.I., and Lust, R.M. (2014). Circadian Influences on Myocardial Infarction. Frontiers in physiology 5.

84.

Durrington, H.J., Farrow, S.N., Loudon, A.S., and Ray, D.W. (2014). The circadian clock and asthma. Thorax 69, 90-92.

25

85.

Kaladchibachi, S.A., Doble, B., Anthopoulos, N., Woodgett, J.R., and Manoukian, A.S. (2007). Glycogen synthase kinase 3, circadian rhythms, and bipolar disorder: a molecular link in the therapeutic action of lithium. Journal of circadian rhythms 5, 3.

86.

Calgaro, S., Boube, M., Cribbs, D.L., and Bourbon, H.M. (2002). The Drosophila gene taranis encodes a novel trithorax group member potentially linked to the cell cycle regulatory apparatus. Genetics 160, 547-560.

87.

Manansala, M.C., Min, S., and Cleary, M.D. (2013). The Drosophila SERTAD protein Taranis determines lineage-specific neural progenitor proliferation patterns. Developmental biology 376, 150-162.

88.

Schuster, K.J., and Smith-Bolton, R.K. (2015). Taranis Protects Regenerating Tissue from Fate Changes Induced by the Wound Response in Drosophila. Dev Cell 34, 119-128.

89.

Chakrabarti, S., Liehl, P., Buchon, N., and Lemaitre, B. (2012). Infectioninduced host translational blockage inhibits immune responses and epithelial renewal in the Drosophila gut. Cell host & microbe 12, 60-70.

90.

Hsu, S.I., Yang, C.M., Sim, K.G., Hentschel, D.M., O'Leary, E., and Bonventre, J.V. (2001). TRIP-Br: a novel family of PHD zinc finger- and bromodomain-interacting proteins that regulate the transcriptional activity of E2F-1/DP-1. The EMBO journal 20, 2273-2285.

91.

Sugimoto, M., Nakamura, T., Ohtani, N., Hampson, L., Hampson, I.N., Shimamoto, A., Furuichi, Y., Okumura, K., Niwa, S., Taya, Y., et al. (1999). Regulation of CDK4 activity by a novel CDK4-binding protein, p34(SEI-1). Genes & development 13, 3027-3033.

26

92.

Liew, C.W., Boucher, J., Cheong, J.K., Vernochet, C., Koh, H.J., Mallol, C., Townsend, K., Langin, D., Kawamori, D., Hu, J., et al. (2013). Ablation of TRIP-Br2, a regulator of fat lipolysis, thermogenesis and oxidative metabolism, prevents diet-induced obesity and insulin resistance. Nat Med 19, 217-226.

93.

Fernandez-Marcos, P.J., Pantoja, C., Gonzalez-Rodriguez, A., Martin, N., Flores, J.M., Valverde, A.M., Hara, E., and Serrano, M. (2010). Normal Proliferation and Tumorigenesis but Impaired Pancreatic Function in Mice Lacking the Cell Cycle Regulator Sei1. PloS one 5, e8744.

94.

Meyer, C.A., Jacobs, H.W., Datar, S.A., Du, W., Edgar, B.A., and Lehner, C.F. (2000). Drosophila Cdk4 is required for normal growth and is dispensable for cell cycle progression. The EMBO journal 19, 4533-4542.

95.

Odajima, J., Wills, Z.P., Ndassa, Y.M., Terunuma, M., Kretschmannova, K., Deeb, T.Z., Geng, Y., Gawrzak, S., Quadros, I.M., Newman, J., et al. (2011). Cyclin E constrains Cdk5 activity to regulate synaptic plasticity and memory formation. Dev Cell 21, 655-668.

96.

Lin, W.-H., He, M., and Baines, R.A. (2015). Seizure suppression through manipulating splicing of a voltage-gated sodium channel. Brain 138, 891-901.

97.

Nagy, Z., and Esiri, M.M. (1998). Neuronal cyclin expression in the hippocampus in temporal lobe epilepsy. Experimental neurology 150, 240247.

27

Chapter 2 ________________________________________________________________ Experimental work

Chapter 2.1 ________________________________________________________________ TARANIS functions with Cyclin A and Cdk1 in a novel arousal center to control sleep in Drosophila (Manuscript published in Current Biology) (2015)

Article

TARANIS Functions with Cyclin A and Cdk1 in a Novel Arousal Center to Control Sleep in Drosophila Highlights tara is a novel sleep-regulatory gene in Drosophila

Authors

d

TARA regulates CycA levels and interacts with CycA to control sleep

Dinis J.S. Afonso, Die Liu, Daniel R. Machado, ..., James E.C. Jepson, Dragana Rogulja, Kyunghee Koh

d

TARA promotes sleep in CycA-expressing PL neurons, a novel arousal center

Correspondence

d

[email protected] d

Cdk1 interacts with tara and CycA and acts in PL neurons to suppress sleep

In Brief The molecular and neural mechanisms of sleep regulation are not well understood. Afonso et al. show that TARANIS promotes sleep by regulating CycA protein levels and inhibiting Cdk1 activity in a novel arousal center.

Afonso et al., 2015, Current Biology 25, 1717–1726 June 29, 2015 ª2015 Elsevier Ltd All rights reserved http://dx.doi.org/10.1016/j.cub.2015.05.037

Current Biology

Article TARANIS Functions with Cyclin A and Cdk1 in a Novel Arousal Center to Control Sleep in Drosophila Dinis J.S. Afonso,1,2,3 Die Liu,1 Daniel R. Machado,1,2,3 Huihui Pan,1 James E.C. Jepson,1,5 Dragana Rogulja,4 and Kyunghee Koh1,* 1Department of Neuroscience, Farber Institute for Neurosciences and Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, PA 19107, USA 2Life and Health Sciences Research Institute (ICVS), School of Health Sciences, University of Minho, 4710-057 Braga, Portugal 3ICVS/3B’s, PT Government Associate Laboratory, 4710-057 Braga/Guimara ˜ es, Portugal 4Department of Neurobiology, Harvard Medical School, Boston, MA 02115, USA 5Present address: UCL Institute of Neurology, London WC1N 3BG, UK *Correspondence: [email protected] http://dx.doi.org/10.1016/j.cub.2015.05.037

SUMMARY

Sleep is an essential and conserved behavior whose regulation at the molecular and anatomical level remains to be elucidated. Here, we identify TARANIS (TARA), a Drosophila homolog of the Trip-Br (SERTAD) family of transcriptional coregulators, as a molecule that is required for normal sleep patterns. Through a forward-genetic screen, we isolated tara as a novel sleep gene associated with a marked reduction in sleep amount. Targeted knockdown of tara suggests that it functions in cholinergic neurons to promote sleep. tara encodes a conserved cell-cycle protein that contains a Cyclin A (CycA)-binding homology domain. TARA regulates CycA protein levels and genetically and physically interacts with CycA to promote sleep. Furthermore, decreased levels of Cyclindependent kinase 1 (Cdk1), a kinase partner of CycA, rescue the short-sleeping phenotype of tara and CycA mutants, while increased Cdk1 activity mimics the tara and CycA phenotypes, suggesting that Cdk1 mediates the role of TARA and CycA in sleep regulation. Finally, we describe a novel wake-promoting role for a cluster of 14 CycA-expressing neurons in the pars lateralis (PL), previously proposed to be analogous to the mammalian hypothalamus. We propose that TARANIS controls sleep amount by regulating CycA protein levels and inhibiting Cdk1 activity in a novel arousal center. INTRODUCTION Most animals sleep, and evidence for the essential nature of this behavior is accumulating [1–3]. However, we are far from understanding how sleep is controlled at a molecular and neural level. The fruit fly, Drosophila melanogaster, has emerged as a powerful model system for understanding complex behaviors such as sleep [4, 5]. Mutations in several Drosophila genes have been

identified that cause significant alterations in sleep [5–13]. Some of these genes were selected as candidates because they were implicated in mammalian sleep [10, 11]. However, others (such as Shaker and CREB) whose role in sleep was first discovered in Drosophila [6, 12] have later been shown to be involved in mammalian sleep [14, 15], validating the use of Drosophila as a model system for sleep research. Since the strength of the Drosophila model system is the relative efficiency of large-scale screens, we and other investigators have conducted unbiased forward-genetic screens to identify novel genes involved in sleep regulation [6–9, 16]. Previous genetic screens for short-sleeping fly mutants have identified genes that affect neuronal excitability [6, 7], protein degradation [9, 16], and cell-cycle progression [8]. However, major gaps remain in our understanding of the molecular and anatomical basis of sleep regulation by these and other genes. Identifying the underlying neural circuits would facilitate the investigation of sleep regulation. The relative simplicity of the Drosophila brain provides an opportunity to dissect these sleep circuits at a level of resolution that would be difficult to achieve in the more complex mammalian brain. Several brain regions, including the mushroom bodies, pars intercerebralis, dorsal fan-shaped body, clock neurons, and subsets of octopaminergic and dopaminergic neurons, have been shown to regulate sleep [17–23]. However, the recent discovery that Cyclin A (CycA) has a sleep-promoting role and is expressed in a small number of neurons distinct from brain regions detailed above [8] suggests the existence of additional neural clusters involved in sleep regulation. From an unbiased forward-genetic screen, we discovered taranis (tara), a mutant that exhibits markedly reduced sleep amount. tara encodes a Drosophila homolog of the Trip-Br (SERTAD) family of mammalian transcriptional coregulators that are known primarily for their role in cell-cycle progression [24–27]. TARA and Trip-Br proteins contain a conserved domain found in several CycA-binding proteins [26]. Our research shows that tara regulates CycA levels and genetically interacts with CycA and its kinase partner Cyclin-dependent kinase 1 (Cdk1) [28] to regulate sleep. Furthermore, we show that a cluster of CycA-expressing neurons in the dorsal brain lies in the pars lateralis (PL), a neurosecretory cluster previously proposed to

Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved 1717

control taras tara1/s

B

C

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A

(s

)

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)

Figure 1. Sleep Phenotypes of tara Mutants

(A) Sleep profile of background control (white circles), taras132 (taras, black X’s), and tara1/s132 (tara1/s, black squares) female flies (n = 50–64) in 1 30-min bins. The white and black bars below 20 the x axis represent 12-hr-light and 12-hr-dark Df(Exel7329) periods, respectively. tara-A tara-B 10 (B) Total daily sleep amount for control and tara l /+ e/+ 1/+ s e/s 1/s female flies of the indicated genotypes (n = 44–72). tro s ATG ATG n tara In this and subsequent figures, s132 and e01264 0 co alleles are referred to as s and e, respectively. (C) Schematic of the genomic region of the tara D E F 3 120 40 locus. Gray dashed lines indicate transposon *** insertion sites. The Exel7329 deficiency removes * 30 2 *** 80 most of tara-A and all of tara-B coding regions as 20 indicated. 1 40 (D–G) Waking activity (activity counts per waking 10 minute) (D), sleep-bout duration (E), sleep-bout 0 0 0 + e/+ /+ s /s 1/s l + + s + s s number (F), and sleep latency (time from lights off + s + + / s s l l / / / / / / / / / / 1 e e 1 e 1 ro s ro s e 1 ro s e 1 to the first sleep bout) (G) for the same female flies nt nt nt tara o tara tara o o c c c shown in (B). Sleep-bout duration is not normally distributed and is shown in simplified box plots, G H I * 1000 where the median and interquartile range are *** 1000 120 represented. *** 800 800 (H) Total daily sleep amount of control and 600 600 80 * Df(3R)Exel7329 female heterozygotes in trans to 400 400 either a wild-type (Df/+) or taras132 (Df/s) allele 40 200 200 (n = 35–102). 0 (I) Total daily sleep of control, taras132, and precise 0 0 l s Pr ol Df/+ Df/s + e/+ /+ s /s 1/s o excision (taraPr) female flies (n = 16–36). l r / r t 1 e n nt tro s o (J) Sleep profile of female flies of the indicated tara tara o n c tara c co genotypes (n = 53–58). The white and black bars below the x axis represent 12-hr-light and 12-hrtara-A tara-B L J K *** *** control dark periods, respectively. 1000 *** *** Ub-tara 1.2 (K) Total daily sleep amount for the same flies e/s 800 Ub-tara; tara 30 showed in (J). ** tarae/s 600 0.8 * (L) tara-A and tara-B mRNA levels relative to actin 400 ns 20 mRNA levels in head extracts of indicated geno0.4 200 types (n = 3–6). For each experiment, relative tara 0 l ra e/s e/s mRNA levels of the control flies were set to 1. 0 o 10 s s r a s e/ 1/s s e/ 1/s nt -t ra ra Mean ± SEM is shown. *p < 0.05, **p < 0.01, ***p < co Ub ta ; ta tara tara 0.001; ns, not significant, one-way ANOVA fola r 0 -ta lowed by Tukey post hoc test (B, D, F, G, K, and L) b U or Dunnett post hoc test relative to control flies (H and I); Kruskal-Wallis test (E). For simplicity, only significant differences between the control and each mutant genotype (above the bar for the mutant) and those between taras132, tarae01264/s132, and tara1/s132mutants (above the line for the mutant pair) are indicated. See also Figure S1. 32

** *** ** ***

P 1 GF e0

4 26

be analogous to the mammalian hypothalamus, a major sleep center [29, 30]. Knockdown of tara and increased Cdk1 activity in CycA-expressing PL neurons, as well as activation of these cells, reduces sleep. Collectively, our data suggest that TARA promotes sleep through its interaction with CycA and Cdk1 in a novel arousal center. RESULTS Identification of tara as a Sleep-Regulatory Gene in Drosophila In an ongoing forward-genetic screen for sleep and circadian mutants in Drosophila [31], we identified a novel transposon insertion line (s132) that resulted in a substantial reduction in daily sleep (Figures 1A, 1B, S1A, and S1B). Sleep was reduced in both female and male mutants relative to background controls. Using inverse-PCR, we mapped the s132 P-element inser-

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tion to the tara locus (Figure 1C), which suggests that TARA has a previously unappreciated role in sleep regulation. The tara transcription unit generates two transcripts (A and B) with alternative transcriptional and translational start sites [26] (http://flybase. org; Figure 1C). The two protein isoforms are identical except for a small number of N-terminal amino acids and appear to be functionally interchangeable [26]. For detailed characterization of the sleep phenotypes of tara mutants, we obtained two additional transposon insertions in the tara locus (tara1 and tarae01264) from Drosophila stock centers (Figure 1C). s132 homozygotes are viable, but tara1 and tarae01264 homozygotes are lethal, suggesting that 1 and e01264 are stronger alleles than s132. Consistent with this view, when combined in trans with s132, the lethal alleles exhibited a greater reduction in sleep than s132 (Figures 1A, 1B, S1A, and S1B). The strong tara alleles resulted in a significant reduction in sleep even as heterozygotes (Figures 1B and

1718 Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved

A

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S1B). Whereas waking activity (activity counts per minute awake) was slightly increased in some tara mutants, it was not increased in strong allelic combinations (Figures 1D and S1C). Sleep-bout duration in both females and males was reduced in strong allelic combinations (Figures 1E and S1D), which suggests that TARA plays a role in sleep maintenance. The number of sleep bouts was markedly reduced in females with strong tara mutations (Figure 1F) and was unchanged in tara males (Figure S1E). In addition, sleep latency (time from lights off to the first sleep bout) was significantly increased in strong tara mutants (Figures 1G and S1F), revealing a role for TARA in sleep initiation. Taken together, our data demonstrate that tara is a novel sleep gene essential for sleep initiation and maintenance. We undertook additional experiments to rule out the possibility that secondary, background mutations are responsible for the sleep phenotype in tara mutants. First, a deficiency line deleting the tara locus did not complement the s132 allele (Figures 1H and S1G). Second, precise excision of the s132 insertion by transposase-mediated mobilization restored normal sleep (Figures 1I and S1H). Third, ubiquitous expression of tara-B [26] restored sleep to nearly normal levels (Figures 1J, 1K, and S1I). These data confirm that disruption of tara is indeed the underlying cause of the severe sleep reduction in tara mutants. As shown in Figures 1B and S1B, three tara allelic combinations (s132, e01264/s132, and 1/s132) yielded varying degrees of sleep reduction, suggesting that tara1 is the strongest allele and taras132 is the weakest. To determine whether differences in tara mRNA levels mediate varying phenotypic strengths, we performed qRT-PCR using primers designed to distinguish between the two tara isoforms. taras132 homozygous mutants had almost no detectable tara-A mRNA and an 50% reduction in tara-B mRNA levels relative to control flies (Figure 1L). Like taras132 mutants, tara1/s132 flies had almost no detectable tara-A mRNA, but tara-B transcripts were further reduced, indicating that tara1 is a null or strongly hypomorphic allele. In tarae01264/s132 flies, tara-A mRNA levels were slightly higher than in tara1/s132 flies while tara-B mRNA levels were lower than in taras132 homozygous flies. These results demonstrate that the amount of daily sleep correlates with tara levels. Collectively, the above data establish tara as a novel sleep regulatory gene.

/s

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Figure 2. Circadian Phenotypes and ClockIndependent Sleep Loss of tara Mutants (A) Representative circadian actogram of individual control and tara1/s132 male flies in DD. Gray and black bars above the actogram indicate subjective day and night, respectively. (B) Cycling of PER protein (green) in small ventral lateral neurons (s-LNvs) is normal in tara1/s132 brains. Samples were dissected at indicated Zeitgeber times (ZT) and stained for PER and PDF (red), which was used to identify s-LNvs. Scale bar, 10 mm. (C) Cycling of PER is also normal in a cluster of dorsal neurons (DN1s). Scale bar, 20 mm. (D) Total daily sleep amount in LL and DD for females of indicated genotypes (n = 32–79 for LL; n = 39–96 for DD). Sleep levels on the third day in constant conditions are shown. Mean ± SEM is shown. ***p < 0.001, Dunnett post hoc tests relative to control flies (D). See also Figure S2.

Sleep Loss in tara Mutants Is Independent of the Circadian Clock and Light To examine whether tara mutants exhibit circadian phenotypes, we monitored their locomotor activity in constant darkness (DD). Most tara1/s132 mutants were arrhythmic or weakly rhythmic and the amplitude of their circadian rhythmicity was reduced, but the period length of all tara mutants was indistinguishable from that of control flies (Figures 2A and S2A). Moreover, daily cycling of the core clock protein PERIOD (PER) in tara1/s132 mutants was similar to that in wild-type controls in two sets of clock neurons (Figures 2B and 2C), which suggests that dampened rhythmicity in these mutants is not due to a defect in the core molecular clock. Since arrhythmicity does not necessarily lead to short sleep (e.g., per and timeless mutants do not have reduced sleep [32]), the rhythm phenotype of tara mutants may not be the cause of the sleep phenotype. Our data showing that tarae01264/s132 mutants displayed almost as severe a sleep reduction as tara1/s132 but were largely rhythmic (Figures 1B, S1B, and S2A) support the view that the sleep and circadian phenotypes in tara mutants may not be linked. To test whether the sleep phenotype in tara mutants was due to arrhythmicity, we assayed sleep in constant light (LL), in which both control and mutant flies are arrhythmic. Indeed, tara mutants had greatly reduced sleep compared with controls in LL, demonstrating that the shortsleeping phenotype is not caused by arrhythmicity (Figures 2D and S2B). The short-sleeping phenotype was also observed in DD (Figures 2D and S2B), suggesting that TARA’s role in sleep is independent of light. Of note, in both LL and DD, tara1/s132 mutants lost over 80% of sleep relative to control flies, which is one of the most severe phenotypes documented among sleep mutants. These data show that tara mutants exhibit a striking reduction in sleep amount, independent of the circadian clock and light conditions. TARA Is Required in Neurons to Control Sleep Levels To examine the spatial requirements for TARA in regulating sleep, we generated a polyclonal antibody against the TARA protein (see Experimental Procedures). In western blots, the antibody recognized a band that is upregulated when TARA is overexpressed in Drosophila S2 cells. As expected, this band was

Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved 1719

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Figure 3. TARA Regulates Sleep in Neurons

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markedly downregulated in head extracts of tara1/s132 mutants compared with those of control flies (Figure 3A). The identity of the band was further examined by western analysis of a previously generated GFP fusion trap in the tara locus (YB0035) [33], which we termed tara::GFP. The GFP exon is located upstream of the common second coding exon of both tara-A and tara-B isoforms (Figure 1C) and is expected to be incorporated into both isoforms close to the N terminus. The presumed TARA band in western blots was shifted by the addition of GFP in head extracts of tara::GFP flies (Figure S3A), which confirms that the band indeed represents the TARA protein. Because the polyclonal antibody did not yield a specific signal when used for immunohistochemistry, we employed the TARA::GFP fusion protein to determine the expression pattern of TARA.

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(A) Western blot showing a marked reduction of TARA in tara1/s132 mutants compared with control flies. Head extracts of control flies and tara mutants (lanes 1 and 2) as well as S2 cell extracts transfected with an empty UAS vector or a UAStara construct under the control of actin-Gal4 (lanes 3 and 4) were probed with a polyclonal antibody to TARA. The band that corresponds to TARA can be readily recognized by the upregulation in S2 cells transfected with tara cDNA. # denotes non-specific labeling or a degradation product of TARA. MAPK was used to control for loading. (B) Maximal-intensity projection of confocal slices of the adult brain showing widespread expression of TARA::GFP. Scale bar, 50 mm. (C) Representative confocal sections of tara::GFP adult brains costained with antibodies to GFP and ELAV or REPO (neuronal or glial marker, respectively). Each panel shows a single confocal slice of a region ventral to the mushroom bodies. Scale bar, 10 mm. (D) Pan-neuronal knockdown of tara markedly reduces sleep. Pan-neuronal elav-Gal4 or nsyb-Gal4 was used to drive a combination of two UAS-tara RNAi constructs and UAS-dcr2 (elav>tara RNAi and nsyb>tara RNAi, respectively). Flies harboring the two UAS-tara RNAi constructs and UAS-dcr2 without a driver (+ > tara RNAi) and those harboring a driver and UAS-dcr2 (elav > + or nsyb > +) served as controls (n = 31–58). (E) Adult-stage expression of tara partially rescues the tara short-sleeping phenotype. Daily sleep is presented for females of the indicated genotypes in the absence (white bar) or presence (black or red bar) of RU486 (n = 24–32). Data from parental control flies show that RU486 by itself did not affect sleep. (F) Knockdown of tara in cholinergic neurons reduces sleep. For each Gal4, total daily sleep of females expressing tara RNAi under the control of the driver (black bar) was compared to parental controls (white bar) (n = 30–173). The sleep phenotype of flies in which tara was knocked down in dopaminergic neurons was not determined due to lethality. Mean ± SEM is shown. ***p < 0.001, Dunnett post hoc test relative to both parental controls (D), t test with Bonferroni correction (E), Tukey post hoc test relative to both parental controls (F). See also Figure S3.

Homozygotes for the tara::GFP allele did not exhibit altered sleep levels or circadian phenotypes (Figures S2A, S3B, and S3C), indicating that the TARA::GFP fusion protein is functional. Since the GFP coding region is inserted into the tara locus in the genome, the TARA::GFP expression pattern is likely to reflect endogenous TARA expression accurately. We thus examined the localization of TARA::GFP in the adult nervous system using an anti-GFP antibody. TARA::GFP was widely expressed throughout the adult brain (Figure 3B). Costaining with neuronal and glial markers (ELAV and REPO, respectively) demonstrated that TARA is expressed in most, perhaps all, neurons but excluded from glial cells (Figure 3C). Given the expression pattern, we sought to demonstrate a role for neuronal TARA in regulating sleep. We used RNAi to

1720 Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved

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Figure 4. tara Interacts with CycA and Regulates CycA Levels (A) Daily sleep for female flies of indicate genotypes demonstrating a synergistic interaction between cycAEY11746/+ and taras132 (n = 43–60). (B) Daily sleep for female flies of the indicated genotypes. Pan-neuronal knockdown of tara by RNAi (elav>tara RNAi) was more effective at suppressing sleep in a CycAC8LR1/+ background than in a control background (n = 46–52). (C) TARA and CycA form a complex in S2 cells. HA-tagged CycA was immunoprecipitated with an anti-HA antibody, and anti-TARA and anti-HA antibodies were used for western blotting. The experiment was repeated three times with similar results. (D) Maximal-intensity confocal projections of the dorsal half of the central brain of representative control and tara1/s132 adult females immunostained with an antibody to CycA. Scale bar, 50 mm. (E) The central brain of an adult fly in which PL-Gal4 was used to express membrane-targeted CD8::GFP. Scale bar, 50 mm. Images on the right show colocalization of CycA and GFP driven by PL-Gal4 in the brain region indicated by the rectangle. (F) The central brain of a fly in which the synaptic marker HA::SYT was expressed using PL-Gal4. The brain was costained with anti-HA and antiCycA. The rectangle indicates the region magnified in the images on the right. Scale bar, 50 mm. Mean ± SEM is shown. *p < 0.05; ***p < 0.001; ns, not significant, Tukey post hoc test. See also Figure S4.

CycA HA

reduce TARA expression specifically in neurons. As expected, driving tara RNAi with pan-neuronal drivers elav-Gal4 and nsyb-Gal4 resulted in a substantial reduction in daily sleep levels (Figures 3D and S3D). Reduced TARA expression by RNAi-mediated knockdown was confirmed by western analysis (Figure S3E). We next examined whether tara functions in the adult fly to regulate sleep by utilizing a UAS site in the s132 insertion to drive tara expression. We used GeneSwitch (GS), an RU486-dependent GAL4 protein that allows temporal control of transgenic expression [34]. Adult specific pan-neuronal expression of tara using the elav-GS driver partially rescued the short-sleeping phenotype of tara mutants (Figure 3E), demonstrating an adult function of tara, although the incomplete adult-stage rescue suggests a potential developmental role as well. To identify neuronal groups where TARA acts to control sleep, we utilized the Gal4/UAS system to target tara RNAi expression to subsets of neurons. We examined several neurotransmitter systems as well as brain regions involved in sleep regulation such as the mushroom bodies (MB), dorsal fan-shaped body (dFSB), pars intercerebralis (PI), and clock cells. Only tara knockdown by Cha-Gal4 produced a significant reduction in sleep (Figure 3F). These data suggest that cholinergic neurons likely mediate the effects of TARA on sleep.

tara Interacts with CycA to Control Sleep and Regulates CycA Levels Since CycA has been shown to promote sleep in Drosophila [8], and since TARA contains a conserved CycA binding homology motif, we tested whether tara and CycA act in a common genetic pathway to regulate sleep. To do so, we generated double mutants and compared their sleep behavior with those of wildtype control and single mutant flies. The CycAEY11746/+ heterozygous mutation did not cause reduced sleep on its own, but it led to a significant reduction in sleep when combined with the taras132 hypomorphic mutation that has a moderate sleep phenotype (Figures 4A and S4A). This interaction was confirmed using a second allele of CycA (CycAC8LR1/+) and tara RNAi (Figure 4B). Further, CycA did not exhibit a genetic interaction with the DATfmn shortsleeping mutant [13] (Figure S4B), demonstrating the specificity of the interaction between tara and CycA. These data reveal a synergistic interaction between tara and CycA and suggest they act in the same pathway to influence sleep. Given the genetic interaction between tara and CycA, the presence of a CycA-binding homology domain in TARA, and the fact that Trip-Br1/2, mammalian homologs of TARA, have been shown to bind CycA [24], we tested whether TARA physically binds CycA in a co-immunoprecipitation assay. Indeed, we found that TARA co-immunoprecipitated with CycA in

Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved 1721

Drosophila S2 cells (Figure 4C), suggesting that they can form a complex. We next asked whether CycA levels are altered in tara mutants. We performed whole-mount immunostaining of adult brains using a CycA antibody previously shown to detect a dorsal set of CycA-positive neurons [8] (another CycA antibody previously used to detect a few additional clusters of CycA-expressing neurons is no longer available). We found that CycA protein levels were greatly reduced in the adult brain of tara mutants (Figure 4D). In contrast, CycA protein levels were not reduced in DATfmn mutants (Figure S4C), which demonstrates the specificity of the regulation of CycA levels by TARA. CycA mRNA levels were not affected in tara mutants (Figure S4D), indicating that TARA regulates CycA levels post-transcriptionally. Our data suggest that TARA promotes sleep in part through regulation of CycA protein levels. We noticed that the dorsal CycA cluster might correspond to the pars lateralis (PL) [35], so we drove expression of CD8::GFP using PL-Gal4, a driver expressed in the PL neurons [36], while simultaneously labeling brains for CycA. Both GFP and CycA were expressed in 14 neurons with large cell bodies in the dorsal brain (Figures 4E and S4E). The striking overlap seen between the GFP and CycA signals demonstrates that the dorsal CycA neurons indeed lie in the PL. This is significant because the PL, along with the pars intercerebralis, shares several features with the mammalian hypothalamus, a major sleep center [29, 30]. However, a possible contribution of the PL to sleep regulation has not been previously explored. We employed the PL driver to determine whether the CycA-expressing cells were present in tara mutants. By examining flies expressing CD8::GFP under the control of PL-Gal4, we confirmed that the PL neurons were indeed present (Figure S4F). Interestingly, CycA protein was observed not only in cell bodies, but also in discrete puncta that appeared to be synapses (Figure 4D). This is noteworthy because according to the synaptic homeostasis hypothesis, waking activity leads to a net increase in synaptic strength, whereas sleep leads to overall downscaling of synapses [37]. To determine whether these puncta represent synapses, we used PL-Gal4 to express a synaptic marker (HA:: SYT) [38] and demonstrated that CycA indeed localized to synaptic regions (Figure 4F). We note that CycA protein levels were downregulated in both cell bodies and synaptic regions in tara mutants (Figure 4D). CycA levels and function at synapses, under the control of TARA, may be important for normal sleep. TARA Regulates Sleep in CycA-Expressing PL Neurons, which Define a New Arousal Center To address whether TARA is required in CycA-expressing cells for sleep regulation, we examined the sleep phenotype of flies in which tara was knocked down using the PL driver. We found that PL-specific tara knockdown significantly reduced sleep (Figure 5A). We note that this manipulation produced a weaker sleep reduction than pan-neuronal knockdown of tara (Figure 3D), which suggests that TARA likely functions in additional groups of neurons to regulate sleep. Our results pointed to a possible role of the PL neurons in sleep regulation. Indeed, we found that activation of the PL neurons via expression of the bacterial sodium channel NaChBac [39] led to decreased sleep (Figures 5B, 5C, S5A, and S5B). In contrast, ac-

tivity levels during waking periods were not affected by PL activation (Figures 5D and S5C). Sleep-bout duration was markedly decreased while sleep-bout number showed little change, and sleep latency was significantly increased in flies with activated PL neurons (Figures 5E–5G, and S5D–S5F). These data suggest that activation of PL neurons promote wakefulness by delaying sleep onset and impairing sleep maintenance. Adult-stage specific activation of these neurons using the warmth-activated cation channel TrpA1 [40] also reduced sleep, demonstrating that this cell cluster functions in adult animals to promote wakefulness (Figures 5H, 5I, and S5G). Further, blocking the activity of PL neurons with tetanus toxin [41] significantly increased sleep (Figure 5J), which confirms the wake-promoting role of these neurons. The above data identify the PL neurons as a novel arousal center and demonstrate that TARA acts, at least in part, in CycA-positive PL neurons to promote sleep. tara and Cdk1 Interact Antagonistically to Regulate Sleep CycA has been shown to bind Cdk1 and can either increase or decrease Cdk1 activity depending on the cellular context [28, 42]. We therefore asked whether Cdk1 also interacts with tara for sleep regulation. We introduced a heterozygous Cdk1GT-000294/+ mutation (the GT-000294 insertion is in the coding region and is likely to be a null allele) into a tara mutant background and compared their sleep with tara and Cdk1 single mutants as well as with wild-type control flies. We found that the Cdk1GT-000294/+ heterozygous mutation did not cause a sleep phenotype in a wild-type background, but it resulted in a substantial rescue of the tara sleep phenotype (Figures 6A and 6B). We confirmed the antagonistic interaction between tara and Cdk1 using a second allele of Cdk1 (Cdk1c03495/+) (Figure S6A). The Cdk1GT-000294/+ heterozygous mutation did not rescue the short-sleeping phenotype of insomniac (inc) mutants (Figure S6B) [9, 16], which demonstrates that the interaction between tara and Cdk1 is not due to additive effects. In contrast, the Cdk1 mutation did rescue the sleep phenotype of heterozygous CycA null mutants (Figure 6C), consistent with a model in which tara and CycA act together to antagonize Cdk1. Transcript levels of Cdk1 were not significantly affected in tara mutants (Figure S6C), suggesting that the interaction between tara and Cdk1 is not likely to be due to transcriptional regulation of Cdk1 by TARA. The antagonistic interaction between tara and Cdk1 suggests that Cdk1 has a previously unrecognized wake-promoting role. To investigate the potential wake-promoting role of Cdk1, we assayed sleep in flies overexpressing wild-type Cdk1 (Cdk1WT). Since activity of Cyclin-dependent kinases is tightly controlled by a number of regulatory molecules [42–44], we also examined flies overexpressing Cdk1-AF, a mutant Cdk1 protein that has elevated kinase activity due to mutations in inhibitory phosphorylation sites [42]. Because overexpression of Cdk1-AF under the control of elav-Gal4 resulted in lethality, we used the RU486 inducible elav-GS to express Cdk1 specifically in the adult stage. Whereas RU486 had little effect on control flies, flies in which Cdk1-AF was expressed under the control of elav-GS exhibited significantly reduced sleep when fed RU486 (Figure 6D), which indicates that increased Cdk1 activity indeed promotes wakefulness. In contrast, overexpression of Cdk1-WT had little effect on sleep (Figure S6C), presumably because

1722 Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved

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Figure 5. TARA Regulates Sleep in CycAExpressing PL Neurons that Define a New Arousal Center (A) Total daily sleep of female flies of the indicated genotypes (n = 32). (B) Sleep profile for female flies expressing the NaChBac sodium channel under the control of PL-Gal4 (PL > NaChBac) and parental controls (n = 48–63). (C–G) Total daily sleep (C), waking activity (D), sleep-bout duration (E), sleep-bout number (F), and latency to sleep after lights off (G) for the flies shown in (B). (H) Sleep profile for female flies expressing TrpA1 under the control of PL-Gal4 (PL > TrpA1) and parental controls (n = 16–32). Flies were monitored at 29 C, which activates the TrpA1 channel, and at 22 C, which inactivates the TrpA1 channel. Sleep profile for the second day at 29 C is omitted for simplicity. (I) Total daily sleep for flies shown in (H). (J) Female flies expressing functional tetanus toxin under the control of PL-Gal4 (PL > TNT-G) exhibited a significant increase in sleep relative to flies expressing inactive tetanus toxin (PL > TNTIMP) or those carrying either form of tetanus toxin transgene without the PL driver (+ > TNT-G and + > TNT-IMP) (n = 30–32). Mean ± SEM is shown. *p < 0.05, **p < 0.01, ***p < 0.001; ns, not significant; Dunnett post hoc test relative to parental controls (A, C, D, F, G, and I) or PL > TNT-G flies (J); Kruskal-Wallis test (E). See also Figure S5.

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that Cdk1 interacts antagonistically with TARA and CycA and acts in PL neurons to promote wakefulness.

P -G MP -G -IM NT T-I TNT T T N TN > T > > + L > PL + P

overexpression of wild-type Cdk1 alone was not sufficient to increase its kinase activity. To examine whether Cdk1 acts in CycA-expressing cells to regulate sleep, we assayed sleep in flies expressing the Cdk1-AF transgene under the control of PL-Gal4. These flies had significantly reduced sleep compared with parental control flies (Figure 6E). These data provide strong evidence for a novel role of Cdk1 in suppressing sleep. Since CycA is expressed in synaptic regions (Figure 4F), we next asked whether Cdk1 colocalizes with CycA at synaptic regions. To address this question, we expressed MYC-tagged wild-type Cdk1 [45] in PL neurons and found that Cdk1::MYC exhibited marked overlap with CycA puncta at synaptic regions (Figure 6F). Although the synaptic localization of Cdk1::MYC could be an artifact of overexpression, the potential colocalization of Cdk1 and CycA at synaptic regions raises the interesting possibility that synaptic Cdk1 activity may be important for maintaining normal sleep amount. Together, our data demonstrate

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DISCUSSION

From an unbiased forward genetic screen, we have identified a novel sleep regulatory gene, tara. Our data demonstrate that TARA interacts with CycA to regulate its levels and promote sleep. We have also identified Cdk1 as a wake-promoting molecule that interacts antagonistically with TARA. Given the fact that TARA regulates CycA levels, the interaction between TARA and Cdk1 may be mediated by CycA. Our finding that Cdk1 and CycA also exhibit an antagonistic interaction supports this view. The previous discovery that CycE sequesters its binding partner Cdk5 to repress its kinase activity in the adult mouse brain [46] points to a potential mechanism, namely that TARA regulates CycA levels, which in turn sequesters and inhibits Cdk1 activity. TARA and its mammalian homologs (the Trip-Br family of proteins) are known for their role in cell-cycle progression [24–27]. However, recent data have shown that Trip-Br2 is involved in lipid and oxidative metabolism in adult mice [47], demonstrating a role beyond cell-cycle control. Other cell-cycle proteins have also been implicated in processes unrelated to the cell cycle. For example, CycE functions in the adult mouse brain to regulate learning

Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved 1723

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(A) Sleep profile for control (white diamonds), Cdk1GT-000294/+ (gray circles), tarae01264/s132 (black squares), and Cdk1GT-000294/+; tarae01264/s132 (red squares) female flies (n = 50–64). (B) Total daily sleep for the flies shown in (A). (C) Total daily sleep of female flies of the indicated genotypes (n = 81–88). (D) Adult-stage specific expression of Cdk1-AF induced by feeding flies food that contain RU486 diluted in ethanol (EtOH) reduced sleep (n = 31–32). (E) Cdk1-AF expression specifically in CycAexpressing PL neurons resulted in reduced sleep (n = 70–75). (F) The central brain of a fly in which UAS-Cdk1myc was expressed using PL-Gal4. The brain was costained with anti-MYC and anti-CycA. The rectangle indicates the region magnified in the images on the right. Scale bar, 50 mm. Mean ± SEM is shown. ***p < 0.001; ns, not significant; Tukey post hoc test (B and C), Dunnett post hoc test relative to all other controls (D); relative to both parental controls (E). See also Figure S6.

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Figure 6. Cdk1 Interacts Antagonistically with tara and CycA, and Increased Cdk1 Activity Suppresses Sleep

CycA MYC

and memory [46]. Based on the finding that CycA and its regulator Rca1 control sleep, it was hypothesized that a network of cell-cycle genes was appropriated for sleep regulation [8]. Our data showing that two additional cell-cycle proteins, TARA and Cdk1, control sleep and wakefulness provide support for that hypothesis. Moreover, the fact that TARA and CycA, factors identified in two independent unbiased genetic screens, interact with each other highlights the importance of a network of cell-cycle genes in sleep regulation. There are two main regulatory mechanisms for sleep: the circadian mechanism that controls the timing of sleep and the homeostatic mechanism that controls the sleep amount [48]. We have shown that TARA has a profound effect on total sleep time. TARA also affects rhythmic locomotor behavior. Since TARA is expressed in clock cells (our unpublished data), whereas CycA is not [8], it is possible that TARA plays a nonCycA dependent role in clock cells to control rhythm strength. Our finding that tara mutants exhibit severely reduced sleep in constant light suggests that the effect of TARA on sleep amount is not linked to its effect on rhythmicity. Instead, TARA may have a role in the sleep homeostatic machinery, which will be examined in our ongoing investigation.

To fully elucidate how sleep is regulated, it is important to identify the unMYC derlying neural circuits. Here, we have shown that activation of the CycA-expressing neurons in the PL suppresses CycA sleep while blocking their activity increases sleep, which establishes them as a novel wake-promoting center. MERGE Importantly, knockdown of tara and increased Cdk1 activity specifically in the PL neurons leads to decreased sleep. A simple hypothesis, consistent with our finding that both activation of PL neurons and increased Cdk1 activity in these neurons suppress sleep is that Cdk1 affects neuronal excitability and synaptic transmission. Interestingly, largescale screens for short-sleeping mutants in fruit flies and zebrafish have identified several channel proteins such as SHAKER, REDEYE, and ETHER-A-GO-GO [6, 49, 50] and channel modulators such as SLEEPLESS and WIDE AWAKE [51, 52]. Thus, it is plausible that Cdk1 regulates sleep by phosphorylating substrates that modulate the function of synaptic ion channels or proteins involved in synaptic vesicle fusion, as has previously been demonstrated for Cdk5 at mammalian synapses [53]. Whereas our data mapped some of TARA’s role in sleep regulation to a small neuronal cluster, the fact that pan-neuronal tara knockdown results in a stronger effect on sleep than specific knockdown in PL neurons suggests that TARA may act in multiple neuronal clusters. PL-specific restoration of TARA expression did not rescue the tara sleep phenotype (data not shown), further implying that the PL cluster may not be the sole anatomical locus for TARA function. Given that CycA is expressed in a few additional clusters [8], TARA may act in all

1724 Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved

CycA-expressing neurons including those not covered by PLGal4. TARA may also act in non-CycA-expressing neurons. Our data demonstrate that tara knockdown using Cha-Gal4 produces as strong an effect on sleep as pan-neuronal knockdown (Figures 3D and 3F). This finding suggests that TARA acts in cholinergic neurons, although we cannot rule out the possibility that the Cha-Gal4 expression pattern includes some noncholinergic cells. Taken together, our data suggest that TARA acts in PL neurons as well as unidentified clusters of cholinergic neurons to regulate sleep. Based on genetic interaction studies, tara has been classified as a member of the trithorax group genes, which typically act as transcriptional coactivators [26, 54]. However, TARA and TripBr1 have been shown to up- or downregulate the activity of E2F1 transcription factor depending on the cellular context, raising the possibility that they also function as transcriptional corepressors [24, 27]. Interestingly, TARA physically interacts with CycA and affects CycA protein levels but not its mRNA expression. These findings suggest a novel non-transcriptional role for TARA, although we cannot rule out an indirect transcriptional mechanism. The hypothesis that TARA plays a non-transcriptional role in regulating CycA levels and Cdk1 activity at the synapse may provide an exciting new avenue for future research.

1. Cirelli, C., and Tononi, G. (2008). Is sleep essential? PLoS Biol. 6, e216. 2. Rechtschaffen, A., Gilliland, M.A., Bergmann, B.M., and Winter, J.B. (1983). Physiological correlates of prolonged sleep deprivation in rats. Science 221, 182–184. 3. Shaw, P.J., Tononi, G., Greenspan, R.J., and Robinson, D.F. (2002). Stress response genes protect against lethal effects of sleep deprivation in Drosophila. Nature 417, 287–291. 4. Cirelli, C. (2009). The genetic and molecular regulation of sleep: from fruit flies to humans. Nat. Rev. Neurosci. 10, 549–560. 5. Crocker, A., and Sehgal, A. (2010). Genetic analysis of sleep. Genes Dev. 24, 1220–1235. 6. Cirelli, C., Bushey, D., Hill, S., Huber, R., Kreber, R., Ganetzky, B., and Tononi, G. (2005). Reduced sleep in Drosophila Shaker mutants. Nature 434, 1087–1092. 7. Koh, K., Joiner, W.J., Wu, M.N., Yue, Z., Smith, C.J., and Sehgal, A. (2008). Identification of SLEEPLESS, a sleep-promoting factor. Science 321, 372–376. 8. Rogulja, D., and Young, M.W. (2012). Control of sleep by cyclin A and its regulator. Science 335, 1617–1621. 9. Stavropoulos, N., and Young, M.W. (2011). insomniac and Cullin-3 regulate sleep and wakefulness in Drosophila. Neuron 72, 964–976. 10. Agosto, J., Choi, J.C., Parisky, K.M., Stilwell, G., Rosbash, M., and Griffith, L.C. (2008). Modulation of GABAA receptor desensitization uncouples sleep onset and maintenance in Drosophila. Nat. Neurosci. 11, 354–359. 11. Yuan, Q., Joiner, W.J., and Sehgal, A. (2006). A sleep-promoting role for the Drosophila serotonin receptor 1A. Curr. Biol. 16, 1051–1062.

EXPERIMENTAL PROCEDURES Details of experimental procedures are available in the online Supplemental Information. SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures and six figures and can be found with this article online at http://dx.doi.org/ 10.1016/j.cub.2015.05.037. AUTHOR CONTRIBUTIONS K.K. conceived the study, and D.J.S.A. and K.K. designed the experiments and analyzed the data. D.J.S.A. performed the experiments with the help of D.L., D.R.M., J.E.C.J., and H.P., and D.R. identified the dorsal CycA-expressing cells as the pars lateralis cluster. The manuscript was written by K.K. and D.J.S.A. with editorial input from D.R. and J.E.C.J.

12. Hendricks, J.C., Williams, J.A., Panckeri, K., Kirk, D., Tello, M., Yin, J.C., and Sehgal, A. (2001). A non-circadian role for cAMP signaling and CREB activity in Drosophila rest homeostasis. Nat. Neurosci. 4, 1108– 1115. 13. Kume, K., Kume, S., Park, S.K., Hirsh, J., and Jackson, F.R. (2005). Dopamine is a regulator of arousal in the fruit fly. J. Neurosci. 25, 7377– 7384. 14. Douglas, C.L., Vyazovskiy, V., Southard, T., Chiu, S.Y., Messing, A., Tononi, G., and Cirelli, C. (2007). Sleep in Kcna2 knockout mice. BMC Biol. 5, 42. 15. Graves, L.A., Hellman, K., Veasey, S., Blendy, J.A., Pack, A.I., and Abel, T. (2003). Genetic evidence for a role of CREB in sustained cortical arousal. J. Neurophysiol. 90, 1152–1159. 16. Pfeiffenberger, C., and Allada, R. (2012). Cul3 and the BTB adaptor insomniac are key regulators of sleep homeostasis and a dopamine arousal pathway in Drosophila. PLoS Genet. 8, e1003003. 17. Joiner, W.J., Crocker, A., White, B.H., and Sehgal, A. (2006). Sleep in Drosophila is regulated by adult mushroom bodies. Nature 441, 757–760.

ACKNOWLEDGMENTS We thank Drs. Shelagh Campbell, Jae Park, Lynn Cooley, Henri-Marc Bourbon, Kazuhiko Kume, and Amita Seghal, the Bloomington Stock Center, National Institute of Genetics, and the Harvard (Exelixis) Stock Center for fly stocks; Dr. Ralf Stanewsky for the PER antibody; Drs. M. Boudinot and Francois Rouyer for the FaasX software; Dr. William Joiner for the Sleeplab software; Andrea Nam and Katelyn Kallas for technical assistance; and Jennifer Wilson, Drs. Amita Sehgal, Mi Shi, James Jaynes, and Angelique Lamaze for comments on the manuscript. This work was supported by a grant from the National Institutes of Health (R01GM088221 to K.K.) and predoctoral fellowships from the Portuguese Foundation for Science and Technology (SFRH/BD/51726/2011 to D.J.S.A. and SFRH/BD/52321/2013 to D.R.M.). Sequencing was performed at the Kimmel Cancer Center Nucleic Acid Facility, which is supported by a grant from the NIH (P30CA56036). Received: January 2, 2015 Revised: April 20, 2015 Accepted: May 19, 2015 Published: June 18, 2015

REFERENCES

18. Pitman, J.L., McGill, J.J., Keegan, K.P., and Allada, R. (2006). A dynamic role for the mushroom bodies in promoting sleep in Drosophila. Nature 441, 753–756. 19. Crocker, A., Shahidullah, M., Levitan, I.B., and Sehgal, A. (2010). Identification of a neural circuit that underlies the effects of octopamine on sleep:wake behavior. Neuron 65, 670–681. 20. Liu, Q., Liu, S., Kodama, L., Driscoll, M.R., and Wu, M.N. (2012). Two dopaminergic neurons signal to the dorsal fan-shaped body to promote wakefulness in Drosophila. Curr. Biol. 22, 2114–2123. 21. Donlea, J.M., Thimgan, M.S., Suzuki, Y., Gottschalk, L., and Shaw, P.J. (2011). Inducing sleep by remote control facilitates memory consolidation in Drosophila. Science 332, 1571–1576. 22. Chung, B.Y., Kilman, V.L., Keath, J.R., Pitman, J.L., and Allada, R. (2009). The GABA(A) receptor RDL acts in peptidergic PDF neurons to promote sleep in Drosophila. Curr. Biol. 19, 386–390. 23. Parisky, K.M., Agosto, J., Pulver, S.R., Shang, Y., Kuklin, E., Hodge, J.J., Kang, K., Liu, X., Garrity, P.A., Rosbash, M., and Griffith, L.C. (2008).

Current Biology 25, 1717–1726, June 29, 2015 ª2015 Elsevier Ltd All rights reserved 1725

PDF cells are a GABA-responsive wake-promoting component of the Drosophila sleep circuit. Neuron 60, 672–682.

oscillators in the fly circadian circuit and induces multiple behavioral periods. J. Neurosci. 26, 479–489.

24. Hsu, S.I., Yang, C.M., Sim, K.G., Hentschel, D.M., O’Leary, E., and Bonventre, J.V. (2001). TRIP-Br: a novel family of PHD zinc finger- and bromodomain-interacting proteins that regulate the transcriptional activity of E2F-1/DP-1. EMBO J. 20, 2273–2285.

40. Hamada, F.N., Rosenzweig, M., Kang, K., Pulver, S.R., Ghezzi, A., Jegla, T.J., and Garrity, P.A. (2008). An internal thermal sensor controlling temperature preference in Drosophila. Nature 454, 217–220.

25. Sim, K.G., Zang, Z., Yang, C.M., Bonventre, J.V., and Hsu, S.I. (2004). TRIP-Br links E2F to novel functions in the regulation of cyclin E expression during cell cycle progression and in the maintenance of genomic stability. Cell Cycle 3, 1296–1304. 26. Calgaro, S., Boube, M., Cribbs, D.L., and Bourbon, H.M. (2002). The Drosophila gene taranis encodes a novel trithorax group member potentially linked to the cell cycle regulatory apparatus. Genetics 160, 547–560. 27. Manansala, M.C., Min, S., and Cleary, M.D. (2013). The Drosophila SERTAD protein Taranis determines lineage-specific neural progenitor proliferation patterns. Dev. Biol. 376, 150–162. 28. Pagano, M., Pepperkok, R., Verde, F., Ansorge, W., and Draetta, G. (1992). Cyclin A is required at two points in the human cell cycle. EMBO J. 11, 961–971.

41. Sweeney, S.T., Broadie, K., Keane, J., Niemann, H., and O’Kane, C.J. (1995). Targeted expression of tetanus toxin light chain in Drosophila specifically eliminates synaptic transmission and causes behavioral defects. Neuron 14, 341–351. 42. Ayeni, J.O., Varadarajan, R., Mukherjee, O., Stuart, D.T., Sprenger, F., Srayko, M., and Campbell, S.D. (2014). Dual phosphorylation of cdk1 coordinates cell proliferation with key developmental processes in Drosophila. Genetics 196, 197–210. 43. Su, S.C., and Tsai, L.H. (2011). Cyclin-dependent kinases in brain development and disease. Annu. Rev. Cell Dev. Biol. 27, 465–491. 44. Suryadinata, R., Sadowski, M., and Sarcevic, B. (2010). Control of cell cycle progression by phosphorylation of cyclin-dependent kinase (CDK) substrates. Biosci. Rep. 30, 243–255.

29. Saper, C.B., Scammell, T.E., and Lu, J. (2005). Hypothalamic regulation of sleep and circadian rhythms. Nature 437, 1257–1263.

45. Meyer, C.A., Jacobs, H.W., Datar, S.A., Du, W., Edgar, B.A., and Lehner, C.F. (2000). Drosophila Cdk4 is required for normal growth and is dispensable for cell cycle progression. EMBO J. 19, 4533–4542.

30. de Velasco, B., Erclik, T., Shy, D., Sclafani, J., Lipshitz, H., McInnes, R., and Hartenstein, V. (2007). Specification and development of the pars intercerebralis and pars lateralis, neuroendocrine command centers in the Drosophila brain. Dev. Biol. 302, 309–323.

46. Odajima, J., Wills, Z.P., Ndassa, Y.M., Terunuma, M., Kretschmannova, K., Deeb, T.Z., Geng, Y., Gawrzak, S., Quadros, I.M., Newman, J., et al. (2011). Cyclin E constrains Cdk5 activity to regulate synaptic plasticity and memory formation. Dev. Cell 21, 655–668.

31. Jepson, J.E., Shahidullah, M., Lamaze, A., Peterson, D., Pan, H., and Koh, K. (2012). dyschronic, a Drosophila homolog of a deaf-blindness gene, regulates circadian output and Slowpoke channels. PLoS Genet. 8, e1002671.

47. Liew, C.W., Boucher, J., Cheong, J.K., Vernochet, C., Koh, H.J., Mallol, C., Townsend, K., Langin, D., Kawamori, D., Hu, J., et al. (2013). Ablation of TRIP-Br2, a regulator of fat lipolysis, thermogenesis and oxidative metabolism, prevents diet-induced obesity and insulin resistance. Nat. Med. 19, 217–226.

32. Hendricks, J.C., Lu, S., Kume, K., Yin, J.C., Yang, Z., and Sehgal, A. (2003). Gender dimorphism in the role of cycle (BMAL1) in rest, rest regulation, and longevity in Drosophila melanogaster. J. Biol. Rhythms 18, 12–25. 33. Quin˜ones-Coello, A.T., Petrella, L.N., Ayers, K., Melillo, A., Mazzalupo, S., Hudson, A.M., Wang, S., Castiblanco, C., Buszczak, M., Hoskins, R.A., and Cooley, L. (2007). Exploring strategies for protein trapping in Drosophila. Genetics 175, 1089–1104. 34. Mao, Z., Roman, G., Zong, L., and Davis, R.L. (2004). Pharmacogenetic rescue in time and space of the rutabaga memory impairment by using Gene-Switch. Proc. Natl. Acad. Sci. USA 101, 198–203. 35. Siga, S. (2003). Anatomy and functions of brain neurosecretory cells in diptera. Microsc. Res. Tech. 62, 114–131. 36. Choi, S.H., Lee, G., Monahan, P., and Park, J.H. (2008). Spatial regulation of Corazonin neuropeptide expression requires multiple cis-acting elements in Drosophila melanogaster. J. Comp. Neurol. 507, 1184–1195. 37. Tononi, G., and Cirelli, C. (2014). Sleep and the price of plasticity: from synaptic and cellular homeostasis to memory consolidation and integration. Neuron 81, 12–34. 38. Robinson, I.M., Ranjan, R., and Schwarz, T.L. (2002). Synaptotagmins I and IV promote transmitter release independently of Ca(2+) binding in the C(2)A domain. Nature 418, 336–340. 39. Nitabach, M.N., Wu, Y., Sheeba, V., Lemon, W.C., Strumbos, J., Zelensky, P.K., White, B.H., and Holmes, T.C. (2006). Electrical hyperexcitation of lateral ventral pacemaker neurons desynchronizes downstream circadian

48. Borbe´ly, A.A., and Achermann, P. (1999). Sleep homeostasis and models of sleep regulation. J. Biol. Rhythms 14, 557–568. 49. Rihel, J., Prober, D.A., Arvanites, A., Lam, K., Zimmerman, S., Jang, S., Haggarty, S.J., Kokel, D., Rubin, L.L., Peterson, R.T., and Schier, A.F. (2010). Zebrafish behavioral profiling links drugs to biological targets and rest/wake regulation. Science 327, 348–351. 50. Shi, M., Yue, Z., Kuryatov, A., Lindstrom, J.M., and Sehgal, A. (2014). Identification of Redeye, a new sleep-regulating protein whose expression is modulated by sleep amount. eLife 3, e01473. 51. Wu, M.N., Joiner, W.J., Dean, T., Yue, Z., Smith, C.J., Chen, D., Hoshi, T., Sehgal, A., and Koh, K. (2010). SLEEPLESS, a Ly-6/neurotoxin family member, regulates the levels, localization and activity of Shaker. Nat. Neurosci. 13, 69–75. 52. Liu, S., Lamaze, A., Liu, Q., Tabuchi, M., Yang, Y., Fowler, M., Bharadwaj, R., Zhang, J., Bedont, J., Blackshaw, S., et al. (2014). WIDE AWAKE mediates the circadian timing of sleep onset. Neuron 82, 151–166. 53. Verstegen, A.M., Tagliatti, E., Lignani, G., Marte, A., Stolero, T., Atias, M., Corradi, A., Valtorta, F., Gitler, D., Onofri, F., et al. (2014). Phosphorylation of synapsin I by cyclin-dependent kinase-5 sets the ratio between the resting and recycling pools of synaptic vesicles at hippocampal synapses. J. Neurosci. 34, 7266–7280. 54. Gutie´rrez, L., Zurita, M., Kennison, J.A., and Va´zquez, M. (2003). The Drosophila trithorax group gene tonalli (tna) interacts genetically with the Brahma remodeling complex and encodes an SP-RING finger protein. Development 130, 343–354.

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Current Biology Supplemental Information

TARANIS Functions with Cyclin A and Cdk1 in a Novel Arousal Center to Control Sleep in Drosophila Dinis J.S. Afonso, Die Liu, Daniel R. Machado, Huihui Pan, James E.C. Jepson, Dragana Rogulja, and Kyunghee Koh

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Figure S1. Sleep Phenotypes and Genetic Analysis of tara: Male Data. (A) Sleep profile of homozygous taras132 (black X’s), transheterozygous tara1/s132 (black squares), and their background control males (white circles) (n=60-73). The white and black bars below the X-axis represent light and dark periods, respectively. (B) Control and tara mutant males of the indicated genotypes (n=41-73). (C) Waking activity, (D) sleep bout duration, (E) sleep bout number, and (F) sleep latency at lights off for the same male flies shown in (B). Sleep bout duration is shown in simplified box plots, where the median and interquartile range are represented. (G) Total daily sleep of control and Df(3R)Exel7329 male heterozygotes in trans to either a wild type (Df/+) or taras132 (Df/s) allele (n=35-70). (H) Total daily sleep of control, taras132, and precise excision male flies (n=15-34). (I) Total daily sleep of male flies of the indicated genotypes (n=35-42). Ubiquitous expression of tara-B (Ub-tara) rescued the tara sleep phenotypes. Mean ± SEM is shown. *p < 0.05, **p < 0.01, ***p < 0.001, one-way ANOVA followed by Dunnett post hoc test (G, H); Tukey post hoc test (B, C, E, F, I); Kruskal-Wallis test (D). For simplicity, only significant differences between each mutant and the control and those between taras132, tarae01264/s132, and tara1/s132 mutants are indicated. Related to Figure 1.

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Figure S3. Characterization of taraGFP Flies. (A) Western blot showing a band shift in homozygous taraGFP lysates. (B) Total daily sleep and (C) waking activity of taraGFP and control females (n=47-64). taraGFP flies did not exhibit sleep abnormalities, suggesting TARA::GFP is functional. (D) Pan-neuronal tara knockdown using two independent Gal4 lines (elav and nsyb) resulted in a significant reduction of sleep in males (n=33-63 except forelav > tara RNAi, for which n=11). (E) Western blot shows a marked reduction of TARA levels in females in which tara was knocked down pan-neuronally (elav > tara RNAi). The experiment was performed three times with similar results. MAPK was used as loading control (A,E). Mean ± SEM is shown. ***p < 0.001, ns: not significant, Student’s t-test (B,C); one-way ANOVA followed by Dunnett post hoc test relative to controls (D). Related to Figure 3.

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Figure S4. Genetic Interaction between tara and CycA. (A) Daily sleep for male flies of the indicated genotypes (n=54-64). (B) Daily sleep for female flies of the indicated genotypes (n=40-56). (C) Maximal-intensity confocal projections of the dorsal half of the central brain of representative control and DATfmn adult flies immunostained with an antibody to CycA. In contrast to tara mutants, CycA levels were not altered in DATfmn mutants. Scale bar: 100 µm. (D) CycA mRNA levels relative to actin mRNA levels were not significantly different between tara1/s132 and control flies (n=3). (E) Confocal projection of a female brain (top) and ventral nerve cord (bottom) expressing CD8::GFP under the control of PL-Gal4. GFP expression was observed in a small number of neurons in the dorsal brain. Scale bar: 100 µm. (F) Confocal projection of a representative control or tarad40/e01264 central brain expressing CD8::GFP under the control of PL-Gal4. The CycA-expressing PL neurons were present and grossly normal in morphology in tara mutants. d40 (d, for short) is an imprecise excision allele with a sleep phenotype similar to s132 (see Experimental Procedures). Scale bar: 100 µm. Mean ± SEM is shown. *p < 0.05, ***p < 0.001, ns: not significant, two-way ANOVA followed by Tukey post hoc test (A,B); Student’s t-test (D). Related to Figure 4.

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Figure S5. Activation of CycA-Expressing PL Neurons Suppresses Sleep. (A) Sleep profile for male flies expressing the NaChBac sodium channel under the control of PL-Gal4 (PL > NaChBac) and parental controls (n=48-64). (B) Daily sleep, (C) waking activity, (D) sleep bout duration, (E) sleep bout number, and (F) sleep latency at ZT12 for the flies shown in (A). (G) Total daily sleep for male flies carrying both UAS-TrpA1 and PL-Gal4 (PL > TrpA1, n=32) relative to parental controls (n=16-32). Flies were monitored at 29°C, which activates the TrpA1 channel, and at 22°C, which inactivates the TrpA1 channel. Mean ± SEM is shown. *p < 0.05, **p < 0.01, and ***p < 0.001, one-way ANOVA followed by Dunnett post hoc test relative to controls (B-C, E-G); Kruskal-Wallis test (D). Related to Figure 5. 51

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Figure S6. tara and Cdk1 interact antagonistically to control sleep. (A) Daily sleep for control, Cdk1c03495/+, tarae01264/s132, and Cdk1c03495/+; tarae02164/s132 females (n=23-32). (B) Daily sleep for male flies of the indicated genotypes (n=26-43). (C) Cdk1 mRNA levels of tara1/s132 mutants were comparable to those of control flies. (D) Adult-stage pan-neuronal overexpression of wild-type Cdk1 has little effect on sleep. Flies were fed either RU486 or vehicle (EtOH) (n=31-32). Mean ± SEM is shown. **p < 0.01, ns: not significant, one-way ANOVA followed by Tukey post hoc test (A, B, D); Student’s t-test (C). Related to Figure 6.

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Supplemental Experimental Procedures Fly stocks Flies were maintained at room temperature (RT) on standard food containing molasses, cornmeal, and yeast. UAS-tara-RNAi (JF01421), tara1, taraDf(3R)Exel7329, CycAEY11746, CycAC8LR1, incf00285, Cdk1GT-000294, Cdk1c03495, UAS-Cdk1-myc, UAS–mCD8-GFP, UASNaChBac, UAS-TNT-G, UAS-TNT-IMP, UAS-dicer2 (dcr2), elav-Gal4, nsyb-Gal4, ChaGal4, VGlut-Gal4, Tdc2-Gal4, c309-Gal4, OK107-Gal4, and UAS-TrpA1 were obtained from the Bloomington Stock Center. 104y-Gal4, Dilp2-Gal4, tim-Gal4, Pdf-Gal4, and elav-GS lines were obtained from Dr. Amita Sehgal’s lab; Dh44-Gal4 from the Vienna Drosophila Resource Center; UAS-tara-RNAi (6889R-1) from the National Institute of Genetics, Japan; tarae01264 from the Harvard Exelixis collection; tara::GFP (tarayb0035) from Dr. Lynn Cooley; Ub-tara and taraEP3463 from Dr. Henri-Marc Bourbon; PL-Gal4 from Dr. Jae Park; UAS-HA::syt from Dr. Thomas Schwarz; DATfmn from Dr. Kazuhiko Kume; and UAS-Cdk1-WT-VFP and UAS-Cdk1-AF-VFP from Dr. Shelagh Campbell. Δ311-249

PL-Gal4 contains a Corazonin promoter fragment (504

) [36]. For improved

efficiency of tara knockdown, two RNAi lines (JF01421 and 6889R-1) were combined with UAS-dcr2. All fly lines were outcrossed to an isogenic background control line (iso31) for at least four generations, except for the UAS-tara-RNAi lines.

Generation of excision lines The s132 insertion maps to 573 bp upstream of the tara-B transcription start site. Mobilization of the P-element in the s132 line using Δ2-3 recombinase produced precise excision lines with normal sleep patterns but failed to produce imprecise excision lines.

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To generate an imprecise excision line, we employed a neighboring P-element EP3463, which maps to ~50bp upstream of the tara-B transcriptional start site (www.flybase.org). By mobilizing the EP3463 insertion using Δ2-3 recombinase, we obtained an imprecise excision line d40, in which a portion of the P element was removed. The d40 line was outcrossed 6 times into the control iso31 background. Homozygous tarad40 as well as transheterozygous tarad40/e01264 flies exhibit reduced sleep.

Sleep and circadian assays Four to seven day old flies entrained to a 12h:12h LD cycle for at least 3 days were individually placed in small glass tubes containing 5% sucrose and 2% agar at 25°C except where noted. For the GS experiment, 500 µM RU486 or vehicle (ethanol) was added to the sucrose-agar food. For experiments involving the warmth-induced TrpA1 channel, flies were raised at RT (~21°C) and entrained in LD at 22°C for at least 3 days before being monitored for 1 day at 22°C to establish a baseline, 2 days at 29°C to activate the TrpA1 channel, and 1 day at 22°C to examine recovery. Data from the first day at 29°C as well as the baseline and recovery day are presented. Activity counts were collected in 1-min bins using Drosophila Activity Monitoring (DAM) System (Trikinetics), and sleep was defined as a period of inactivity lasting at least 5 min [S1]. Sleep parameters were analyzed using the PySolo software [S2], except for sleep latency which was analyzed using Sleeplab (William Joiner). Circadian assays were performed essentially as described [31]. Briefly, male flies were monitored in DD using the DAM system (Trikinetics) for six days after being entrained to an LD cycle for at least 3 days. Activity counts collected in 30-min bins were

54

analyzed using the FaasX software (M. Boudinot and F. Rouyer). The software uses χ2 analysis to calculate period length and rhythm power. Rhythm power was determined for all flies including arrhythmic ones, whereas circadian period was determined only for rhythmic flies. Actograms were generated using ClockLab (Actimetrics).

Quantitative real-time reverse-transcriptase PCR (qPCR) Total RNA was extracted from 20-30 female fly heads using TRIzol (Life Technologies), and cDNA was generated using High Capacity cDNA Reverse Transcriptase Kit (Applied Biosystems). qPCR was performed using SYBR green (Applied Biosystems). The following primers were used: 5’-GAA AAA GGC GCC AAA CTT AAA TTA-3’ and 5’-TCG CGG AAT TCA CAT TGG AT-3’ for tara-A; 5’-GAA AAT GTG CAC TGA GGT GAA T-3’ and 5’-GTG TTG GCA TCC TTG CTG T-3’ for tara-B; 5’-GCG GAT GAC ATA AGT GAT GG-3’ and 5’-CAT GAC GCT GTA TAT TTC CGA-3’ for CycA; and 5’-ATG GCG TGG TGT ATA AGG GT-3’ and 5’-AAA TTT CTC TGA TCG CGG TT-3’ for Cdk1.

Antibody production and Western analysis For antigen production, a PCR product coding for 167 amino acids of TARA common to both isoforms was subcloned into the pET-28a protein expression vector using the following primers: 5’-ATG AAT TCT CGC CAT CGG AGC C-3’ and 5’-ATC TCG AGA TGC GGT ACA AAG GGA TG. The His-tagged protein was purified at the Wistar Institute Protein Expression Facility, and injected into rats to generate polyclonal antibody TJR51 (Cocalico Biologicals). TJR51 recognized a ~130kD that was markedly

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reduced in tara mutants. Since the TARA protein is predicted to be smaller at ~100kD, we examined the possibility that the annotated tara transcripts are incomplete. To this end, we generated a full-length UAS-tara-B construct, which contains the entire coding region of the tara-B isoform as annotated in FlyBase (www.flybase.org) but does not contain any intronic or untranslated regions. We transfected this UAS-tara-B construct along with an actin-Gal4 construct in Drosophila S2 cells. Overexpression of the tara-B transcripts in S2 cells resulted in upregulation of the ~130kD band, demonstrating that the band corresponds to the predicted TARA-B protein. As the two TARA isoforms (A and B) differ in size by only 4 amino acids, the single band likely represents both isoforms. Western blot analysis of head extracts and quantification of immunoreactive bands were performed essentially as described [51]. Anti-TARA antibody (TJR51) was used at 1:750, anti-HA (Covance) at 1:1000, and anti-MAPK (Sigma) at 1:10000.

Transient transfection and co-immunoprecipitation (co-IP) For Western analysis, Drosophila S2 cells were transfected with indicated DNA constructs in 24-well plates (150 ng of total DNA) using Effectene (Qiagen). For co-IP experiments, S2 cells were transfected with various combinations of UAS-tara (200 ng) and UAS-HA-CycA constructs [S3] (150 ng) along with actin-Gal4 (100 ng) in 6-well plates using Effectene (Qiagen). UAS vector DNA was included in some conditions to make the total amount of DNA equal in all conditions. Transfected cells were kept at 25°C for 2 days before being harvested. Co-IP was performed essentially as described [51] except that an antibody to HA (Covance) was used for IP and cells were lysed in

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extraction buffer containing 150 mM KCl, 50 mM Tris-Cl at pH 7.0, 10 mM EDTA, 0.2% Triton X-100, 10 mM DTT, and protease inhibitor cocktail (Roche).

Immunohistochemistry Dissected brains were fixed in 4% paraformaldehyde for 30 min at RT. Samples were blocked in 5% normal chicken serum for experiments involving CycA staining, 1% BSA for PER staining, and 5% normal goat serum for other antibody staining. The following primary antibodies were used: rabbit anti-PER [S4] at 1:8000, mouse anti-PDF (DSHB) at 1:2000, rabbit anti-GFP (Invitrogen) at 1:500, goat anti-CycA (Santa Cruz Biotechnology, #15869) at 1:50, anti-ELAV (DSHB) at 1:200, anti-REPO (DSHB) at 1:100, anti-HA (Covance) at 1:1000, and anti-MYC (Invitrogen) at 1:1000. The secondary antibodies, Alexa Fluor 647 goat anti-rabbit, Alexa Fluor 555 goat anti-mouse, and Alexa Fluor 647 chicken anti-goat (Invitrogen) were used at 1:400. Primary and secondary antibodies were incubated at 4°C overnight. Images were obtained on an Olympus Fluoview confocal microscope.

Statistical Analysis Data sets with two groups were compared using t tests. For multiple pairwise comparisons in a data set, t tests with Bonferroni correction were used. For comparisons of multiple groups, one-way ANOVA tests were performed, followed by Dunnett or Tukey post-hoc tests. For genetic interaction experiments, two-way ANOVA tests were performed to test for the interaction. For comparisons of non-normally distributed data,

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Kruskal-Wallis tests were performed, followed by Dunn’s post hoc tests with Bonferroni correction.

Supplemental References S1.

Huber, R., Hill, S.L., Holladay, C., Biesiadecki, M., Tononi, G., and Cirelli, C. (2004). Sleep homeostasis in Drosophila melanogaster. Sleep 27, 628-639.

S2.

Gilestro, G.F., and Cirelli, C. (2009). pySolo: a complete suite for sleep analysis in Drosophila. Bioinformatics 25, 1466-1467.

S3.

Dienemann, A., and Sprenger, F. (2004). Requirements of cyclin a for mitosis are independent of its subcellular localization. Curr Biol 14, 1117-1123.

S4.

Stanewsky, R., Frisch, B., Brandes, C., Hamblen-Coyle, M.J., Rosbash, M., and Hall, J.C. (1997). Temporal and spatial expression patterns of transgenes containing increasing amounts of the Drosophila clock gene period and a lacZ reporter: mapping elements of the PER protein involved in circadian cycling. J Neurosci 17, 676-696.

 

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Chapter 2.2 ___________________________________________________________________ TARANIS regulates circadian rhythms, nocturnal activity, and neuronal morphology Dinis J.S. Afonso and Kyunghee Koh

Title: TARANIS regulates circadian rhythms, nocturnal activity, and neuronal morphology

Dinis J.S. Afonso1,2,3 and Kyunghee Koh1* 1

Department of Neuroscience, the Farber Institute for Neurosciences, and Kimmel

Cancer Center, Thomas Jefferson University, Philadelphia, USA 2

Life and Health Sciences Research Institute (ICVS), School of Health Sciences,

University of Minho, 4710-057 Braga, Portugal 3

ICVS/3B’s, PT Government Associate Laboratory, 4710-057 Braga/Guimarães,

Portugal

Correspondence should be addressed to K.K. ([email protected]).

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Summary

Daily cycling of light and temperature modulates life on Earth. Most organisms have evolved internal molecular clocks to timely anticipate these recurring changes. The core molecular oscillator is composed of interconnected feedback loops that cycle daily in accord with Earth’s rotation. Previously we found that tara interacts with CycA and Cdk1 to regulate sleep and that tara mutants exhibit arrhythmic locomotor behavior in constant darkness (DD). However, the mechanisms through which TARA regulate circadian behavior is unknown. Here, we show that TARA regulates the speed of the molecular oscillator and the neuropeptide output of the pacemaker neurons. We observe that the pace of PERIOD (PER) oscillations is faster in tara mutants and slower in flies overexpressing TARA in the small ventral lateral neurons (sLNvs). Additionally, levels of the neuropeptide PIGMENT DISPERSING FACTOR (PDF), which synchronizes the clock cell network, are markedly decreased in tara mutants. Similarly to Clock (Clk) mutants, tara mutants display a nocturnal pattern of activity in light:dark conditions (LD). Furthermore, tara knockdown or overexpression in sLNvs alters the morphology of their dorsal projection. Taken together, our data suggest that tara functions in multiple steps that link molecular cycling to overt circadian locomotor behavior. This is an ongoing project and future research will clarify how tara regulates locomotor rhythms, nocturnal activity, and neuronal morphology.

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Introduction

The repertoire of behaviors of an animal should be timed and optimized for recurring changes in environmental conditions to promote its survival. Molecular clocks have evolved and are present in different organisms ranging from bacteria to humans [1]. The discovery of the per gene in Drosophila became the foundation to the current knowledge of the molecular mechanisms underpinning circadian rhythms [2]. The core molecular oscillator is composed of transcriptional negative feedback loops where the transcriptional activators CLK and CYCLE (CYC) are negatively regulated by their transcriptional targets per and timeless (tim) [1]. The neural substrates of molecular oscillations are well described in the fly brain. The PER protein is expressed in ~150 clock neurons classified in different clusters based on their anatomy, location and physiology [1]. These 150 neurons can be divided into bilateral and symmetric clusters including the dorsal lateral neurons (LNds), the dorsal neurons (DN1s, DN2s, and DN3s), the lateral posterior neurons (LPNs), the sLNvs, and the large ventral lateral neurons (lLNvs). The sLNvs are believed to be necessary and sufficient to drive circadian locomotor rhythms in nonrecurring environmental conditions (constant darkness and temperature) [3, 4]. In LD cycles, the sLNvs (“morning cells”) drive the peak of activity at dawn, while the 5th sLNv and the LNds (“evening cells”) drive the peak of activity at dusk [3, 4]. The circadian mechanism includes three components that ensure proper interactions between the living organism and the environment: the input component that communicates external light and temperature conditions to the molecular clock [5]; the core molecular clock itself, which is composed of oscillatory changes in the mRNAs and proteins of core clock genes; and the output components that transduce

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molecular oscillations to overt behaviors or physiologic processes. Circadian proteins such as PER and TIM can be used to address where, in the circadian mechanism, a new mutant is acting. If a new mutant is arrhythmic, but exhibits normal PER cycling, we can conclude that the output arm of the circadian mechanism is affected. But, if the PER protein does not cycle or has a different oscillatory pace, the core molecular clock machinery is impacted. Furthermore, a mutant with defects in the input pathway will show anomalous phase response curves and altered TIM degradation in response to light [6], or asynchronous cycling of the PER protein within the clock cells. Here, we describe tara as a novel circadian regulatory molecule. tara was previously described as a regulator of transcription and is the Drosophila homolog of the TRIP-Br (Transcriptional Regulator Interacting with the PHD-Bromodomain) family of proteins [7, 8]. tara mutants have reduced robustness in rest:activity locomotor patterns [9] and tara knockdown restricted to the clock cell network reduces rhythm strength. In addition, our data demonstrate that tara negatively regulates the intrinsic speed of the molecular clock machinery and transcriptionally and post-transcriptionally regulates Pdf, the main synchronizing molecule of the clock cell network. tara not only regulates diurnal behavior and acute responses to light, but is also necessary within the PDF+ cells to modulate the structural morphology of sLNv dorsal projection. In summary, our data suggest that TARA plays important roles in circadian locomotor behavior, nocturnal activity and neuronal morphology.

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Results

tara mutants display arrhythmic locomotor activity

Sleep characterization in tara mutants revealed that the sleep defects were independent of the circadian mechanism [9]. However, tara mutants displayed a dramatic decrease in the robustness of daily oscillations of locomotor circadian activity [9]. These results suggest that tara has a role in the regulation of circadian rhythms in addition to its role in sleep. Pan-neuronal knockdown of tara by RNAi decreased rhythm strength (Figure 1A, B, and E), recapitulating the tara mutant phenotype [9]. To further confirm that tara regulates circadian rhythms, we examined tara mutant flies expressing a tara transgene ubiquitously [7, 9]. Ubiquitous expression of a tara transgene led to a complete rescue of the arrhythmic locomotor phenotype of tara mutants (Figure 1C, D, and E). Interestingly, in contrast to tara mutants or tara knockdown flies whose circadian period length did not significantly differ from that of control flies, the rescued flies exhibited an increased period length (Figure 1E), suggesting that tara may play a role in setting the speed of the core molecular oscillator.

Role of tara in the circadian rhythm of locomotor activity maps to the clock cell network

To assess whether tara functions in clock cells to control rhythmic behavior, we turned to tim-Gal4, which is expressed in all clock cells [10]. tara knockdown in neurons labeled by tim-Gal4 reduced rhythm strength by ~50% compared to sibling

65

controls (Figure 2A, B, and E). Rhythm strength was reduced to a lesser extent using tim-Gal4 than using elav-Gal4, suggesting that tara may also play additional roles in other neurons outside of the clock cell network. Next we reduced tara expression specifically in the LNvs using Pdf-Gal4. tara knockdown with Pdf-Gal4 did not reduce rhythm strength in comparison to sibling controls (Figure 2C, D, and E), which suggests that the role of tara in circadian rhythms is not restricted to the LNvs. Alternatively, Pdf-Gal4 may be a weaker driver than tim-Gal4. Neither tim-Gal4 nor Pdf-Gal4 produced an observable period length phenotype when used to drive tara RNAi (Figure 2E). Although our data show that tara is necessary in clock cells to regulate circadian locomotor activity, tara may also play a role downstream of the clock cells in the output component of the circadian mechanism.

tara overexpression in PDF+ neurons increases the period length of circadian locomotor activity

To further characterize tara function in the molecular clock we next performed overexpression experiments. To this end, we generated a new transgenic fly line containing the previously described UAS-tara FL construct encoding the entire sequence of the B-isoform [9]. The two tara isoforms, A and B, are described to be functionally interchangeable [7]. To overexpress tara, we made use of clock cell specific drivers. tara overexpression with tim-Gal4 lead to lethality precluding behavioral analysis of adult flies. As described above, ubiquitous rescue restored rhythm strength and also increased period length, suggesting that tara may impact the intrinsic speed of the molecular machinery. To address this possibility we turned to Pdf-Gal4 to drive the expression of UAS-tara FL just in PDF+ neurons. Consistent

66

with the ubiquitous rescue, cell-specific tara overexpression increased the period length by ~1.5 hours compared to parental controls without significantly affecting rhythm strength (Figure 3A, B, C, and D). Because the pace of overt locomotor behavior depends on the integrated function of the entire clock cell network [11], driving the expression of tara FL or tara RNAi transgenes with Pdf-Gal4 in flies with a functional clock restricted to the LNvs (e.g., by expressing a per transgene only in LNvs in a per null mutant background) may clarify the effect of TARA levels on period length.

tara modulates the pace of cycling of the core clock component PER

Previously, we showed that the core clock protein PER cycles normally in tara mutants in LD conditions [9]. To further examine the integrity and speed of the molecular clock in tara mutants we looked at PER expression on the 3rd day in DD. We observed robust PER protein cycling even in the strongest tara mutants in both the sLNvs and DN1s, which suggests that the arrhythmic phenotype of tara mutants maps to signaling pathways downstream of the molecular clock (Figure 4A and C). To detect subtle changes in the pace of the molecular clock, we quantified PER cycling in sLNvs. Interestingly, our quantification analysis of PER cycling in sLNvs revealed that the pace of molecular clock is sped up in tara mutants, which is consistent with the lengthened period in locomotor activity of flies overexpressing tara in LNvs. In our previous work, the PER cycling in tara mutants was similar to that of control flies in LD conditions [9]. The modest increase in the speed of the pace of the molecular clock detected on the 3rd day in DD may be due to an accumulation of small effects over three days.

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The increase in the pace of PER cycling in tara mutants and the lengthened locomotor activity rhythm in flies overexpressing tara FL in LNvs suggest that tara is a negative regulator of the intrinsic speed of the core molecular clock machinery. To further test this hypothesis, we quantified PER cycling in flies overexpressing tara in LNvs after three days in constant darkness conditions. Consistently with earlier results, our data demonstrate PER cycling was delayed in TARA overexpressing flies (Figure 5A and B). To address whether the function of TARA in the control of the intrinsic speed of the molecular clock could be due to changes in its levels we quantified TARA::GFP in sLNvs. Our results show that TARA::GFP does not cycle in sLNvs (Figure S1A and B), suggesting that its circadian function likely derives from other aspects rather than circadian changes in its levels. Taken together, these results suggest that tara is a negative regulator of the intrinsic speed of the molecular clock.

Daily oscillations and levels of PDF are reduced in tara mutants

The clock neurons form an interconnected network that gives robustness to behavioral locomotor rhythms [11], and PDF functions as a synchronizing factor within the clock cell network [12, 13]. PDF is expressed in just 16-18 out of the ~150 clock cells per Drosophila brain, but its receptor is expressed in ~60% of all clock cells [14]. Intriguingly, PDF levels in the sLNv dorsal projection have a circadian profile, although rhythmic PDF expression may not be required for circadian locomotor activity [15]. In tara mutants, PDF levels in the sLNv dorsal projection were reduced by ~70% (Figure 4C and Figure 6A and B), suggesting that TARA plays a role in

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regulating PDF levels. To address how tara regulates PDF, we quantified Pdf mRNA from whole heads extracts of tara mutants. In addition to the observed reduction in the PDF protein levels, we found that Pdf mRNA from whole heads extracts was also reduced by ~25%, suggesting that tara regulates Pdf transcription (Figure 6C). However, given the greater reduction in the PDF protein levels compared with the Pdf transcript levels, tara likely regulates PDF at the post-transcriptional level as well. Pdf null mutants are arrhythmic after a few days in DD, and in LD conditions their morning anticipation is absent while their evening peak of activity is advanced [16]. In LD conditions, tara mutants show a morning anticipation and their evening peak of activity is not advanced (Figure 6D). However, it is important to note that PDF is not completely abolished in tara mutants (Figure 6A and B), which suggests that the low levels of PDF are sufficient to drive morning and evening peaks of activity. Nevertheless, the arrhythmic locomotor behavior in tara mutants is likely in part due to low levels of PDF and desynchronization of the clock cell network.

tara mutants display nocturnal behavior in LD conditions

Previous chromatin immunoprecipitation experiments revealed that the core clock transcription factor CLK rhythmically binds to the tara genomic locus [17], suggesting that tara is a direct target of CLK and perhaps mediates some of the Clk mutant phenotypes. Clkjrk mutants have increased activity at night [18, 19], and Clk has a non-circadian role in determining the Drosophila diurnal pattern of activity [18]. Interestingly, tara mutants appear to have a switch in their daily activity profile similar to that of Clkjrk mutants, with increased locomotor activity at night (Figure 6D). In

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Clkjrk mutants, CRYPTOCHROME (CRY) and dopamine levels are elevated [20], suggesting that tara mutants may also have high levels of CRY and enhanced dopamine signaling, which will be interesting to study in the future. In the standard LD conditions, the morning peak of activity has two components: an anticipatory component driven by the circadian clock and a startle response to light onset characterized by a rapid and transient increase in locomotor activity [21]. In several arrhythmic mutants, such as per and tim, startle responses are still observed, but in Clkjrk mutants the startle responses to light onset is absent. Interestingly, tara mutants phenocopy Clkjrk mutants and lack the startle response (Figure 6D). This observation suggests that tara may work with Clk not only to regulate diurnal activity but also to regulate acute responses to light.

tara modulates the structural morphology of the sLNv dorsal projection

The sLNv dorsal projection undergoes daily morphological changes [22-24]. Early in the morning in LD cycles, the sLNv dorsal projection has an “open” conformation while at the end of the day it has a “closed” conformation. Our PDF immunostaining data suggest that the sLNv dorsal projection may be structurally altered, but due to the severe reduction in PDF levels, it was difficult to determine the morphology of the sLNv projection accurately. To test whether tara has a role in the circadian remodeling of sLNv dorsal projection, we used a membrane targeted version of GFP (mCD8::GFP) to label the entire structure of the sLNv dorsal projection, while using Pdf-Gal4 to drive tara RNAi or tara FL specifically in PDF positive cells. In the early morning (ZT3), overexpression of tara FL increased the length and branching of the sLNv dorsal projection, suggesting that tara may promote

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the expansion of the sLNv dorsal projection (Figure 7A, B, and C). Consistent with this interpretation, tara knockdown in PDF+ neurons blunts the length and the branching of the sLNv dorsal projection (Figure 7A, B, and C). Additional work is required to determine whether these are developmental defects or whether TARA has an adult role in the sLNv dorsal projection remodeling. tara knockdown, specifically in PDF+ neurons did not alter rhythm strength (Figure 2C, D, and E), which suggests that the role of tara in the morphology of the sLNv dorsal projection by itself does not cause dampened locomotor rhythms. However, it is possible that altering the structure of the entire network of clock neurons has a cumulative effect on the rhythm strength, as observed with tara knockdown with tim-Gal4.

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Discussion

We have previously shown that most severe tara mutants are arrhythmic in DD, suggesting that tara regulates circadian locomotor activity in addition to sleep [9]. Here, we provide insights into the molecular and neural mechanisms underlying the regulation of locomotor rhythms by TARA. Knockdown and rescue experiments confirmed that TARA is responsible for the arrhythmic phenotype observed in tara mutants. Importantly, tara knockdown in all clock cells using tim-Gal4 reduced rhythm strength, which suggests that TARA acts in clock neurons to regulate locomotor rhythms. Interestingly, our data also demonstrate that TARA acts as negative regulator of the intrinsic speed of the molecular clock and controls the levels of PDF. Lack of tara switches daily patterns of activity so that flies start to behave as nocturnal organisms. Furthermore, tara has a dramatic role in the morphology of the sLNv dorsal projection. Altogether, the evidence points for multiple roles of TARA in controlling circadian rhythms, diurnal patterns of activity, and neuronal morphology. As described above, tara is a novel negative regulator of the intrinsic speed of the molecular clock. Intriguingly, TARA in itself does not appear to cycle in sLNvs. How is TARA regulating the circadian oscillatory pace of PER without cycling itself? Recent work has shown that TRIP-Br1, one of the tara mammalian homologs, forms a protein complex with Protein Phosphatase 2A (PP2A) [25]. In flies PP2A regulates the circadian oscillation of PER [26], and two of the PP2A regulatory subunits: twins and widerborst are expressed in a circadian fashion. It is possible that PP2A regulates TARA phosphorylation in flies as well, and may be the regulatory factor that adds a circadian component to the function of TARA.

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In flies, PP2A antagonizes GLYCOGEN SYNTHASE KINASE 3β (GSK3β)/SHAGGY (SGG) to regulate active zone development at the NMJ [27]. This suggests that tara may interact with PP2A and SGG to regulate synaptic morphology in the adult fly brain. It will be interesting to address whether TARA interacts with PP2A and SGG to modulate the morphology of the sLNv dorsal projection. In longer photoperiodic conditions (summer days), PDF is responsible for separating the peaks of crepuscular activity [28], contributing to the adaptation of organismal physiology to seasonal changes in the duration of light and temperature cycles. Previous work has also demonstrated that, in the absence of CRY, PDF signaling is required for the maintenance of the evening peak of activity [29]. Despite the low levels of PDF observed in tara mutants, their morning and evening peaks of activity do not differ from those observed in control flies. In future work we will search for other behavioral consequences of low PDF levels in tara mutants, including whether tara mutants can adapt to longer photoperiodic conditions and maintain the evening peak of activity in a cry null background. Previous work has found that tara is a direct target of the core molecular clock component CLK [17]. Interestingly, tara phenocopies the increased nighttime activity of Clkjrk flies [30], suggesting that Clk and tara act in the same genetic pathway to regulate light-induced arousal. Previous research has demonstrated that increased CRY activity and dopamine signaling mediate the nocturnal activity of Clkjrk mutants [31]. Thus, we will test whether blocking dopamine signaling and CRY activity restores diurnal activity of tara mutants. Whereas Clk and tara mutants exhibit similar nighttime activity phenotypes, they appear to have opposite circadian period phenotypes; Clkjrk heterozygotes exhibit a long period phenotype in DD conditions [32], while tara mutants have a shorter PER oscillatory pace. This suggests that CLK

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and TARA interact in complex ways for the control of multiple aspects of sleep:wake cycles. From an unbiased forward-genetic screen, we recently discovered tara as a new sleep regulatory molecule [9]. The role of tara in sleep maps in part to a novel arousal center of CycA expressing neurons. Whereas knockdown of tara in PL neurons results in reduced sleep without arrhythmicity, knockdown of tara in clock neurons reduces locomotor activity robustness without reducing sleep amount, suggesting that TARA acts in distinct neural clusters to control sleep amount and circadian locomotor rhythms. Further investigation of tara function will likely yield important insights into the molecular and neural mechanisms of sleep and circadian rhythms.

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Materials and methods

Fly stocks

Flies were maintained at room temperature (RT) on standard food containing molasses, cornmeal, and yeast. tara1, UAS-tara-RNAi (JF01421), UAS-dicer2 (dcr2), UAS–mCD8-GFP, and elav-Gal4 were obtained from the Bloomington Stock Center. UAS-tara-RNAi (6889R-1) was obtained from the National Institute of Genetics, Japan; tarae01264 from the Harvard Exelixis collection; Ub-tara from Dr. Henri-Marc Bourbon. For improved efficiency of tara knockdown, two RNAi lines (JF01421 and 6889R-1) were combined with UAS-dcr2. All fly lines were outcrossed to an isogenic background control line (iso31) for at least five generations with exception for the UAS-tara-RNAi lines that were not backcrossed.

Circadian assays

Four to seven day old flies entrained to a 12h:12h LD cycle for at least 3 days were individually placed in small glass tubes containing 5% sucrose and 2% agar at 25°C except where noted. Activity counts were collected in 1-min bins using Drosophila Activity Monitoring (DAM) System (Trikinetics). Circadian assays were performed essentially as described [33]. Briefly, male and female flies were monitored in DD using the DAM system (Trikinetics) for six days after being entrained to an LD cycle for at least 3 days. Activity counts collected in 30-min bins were analyzed using the FaasX software (M. Boudinot and F. Rouyer, Institut de Neurobiologie Alfred Fessard, CNRS, France). The software uses χ2

75

analysis to calculate period length and rhythm power. Rhythm power was determined for all flies including arrhythmic ones, whereas circadian period was determined only for rhythmic flies. Morning and evening anticipation analysis was also performed with FaasX software. Actograms were generated using ClockLab (Actimetrics).

Immunohistochemistry

Dissected brains were fixed in 4% paraformaldehyde for 30 min at RT. Samples were blocked in 1% bovine serum albumin (BSA); primary and secondary antibodies were diluted in 0.1% BSA. The following primary antibodies were used: anti-PER at 1:8000 (rabbit 13.1), mousse anti-PDF at 1:2000 (mouse C7s) from Developmental Studies Hybridoma Bank (Iowa City, USA), and rabbit anti-GFP (Invitrogen) at 1:500. The secondary antibodies, Alexa Fluor 647 goat anti-rabbit and Alexa Fluor 488 goat anti-mouse (Invitrogen) were used at 1:500. Primary and secondary antibodies were incubated at 4°C overnight. Images were obtained on an Olympus Fluoview confocal microscope.

Statistical Analysis

Data sets with two groups were compared using t tests. For comparisons of multiple groups, one-way ANOVA tests were performed, followed by Dunnett or Tukey post-hoc tests. For genetic interaction experiments, two-way ANOVA tests were performed to test for the interaction.

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Acknowledgments

We thank Drs. Shelagh Campbell, Jae Park, Lynn Cooley, and Henri-Marc Bourbon, the Bloomington Stock Center, National Institute of Genetics, and the Harvard (Exelixis) Stock Center for fly stocks; Drs. M. Boudinot and François Rouyer for the FaasX software; Hui Hui Pan, Andrea Nam, and Katelyn Kallas for technical assistance, and Alexandra Kenny for editorial input.

Author contributions

D.J.S.A. and K.K. conceived the study, designed the experiments and analyzed the data. D.J.S.A. performed the experiments. The manuscript was written by D.J.S.A. and K.K.

77

References

1.

Allada, R., and Chung, B.Y. (2010). Circadian organization of behavior and physiology in Drosophila. Annu Rev Physiol 72, 605-624.

2.

Konopka, R.J., and Benzer, S. (1971). Clock mutants of Drosophila melanogaster. Proceedings of the National Academy of Sciences of the United States of America 68, 2112-2116.

3.

Grima, B., Chelot, E., Xia, R., and Rouyer, F. (2004). Morning and evening peaks of activity rely on different clock neurons of the Drosophila brain. Nature 431, 869-873.

4.

Stoleru, D., Peng, Y., Agosto, J., and Rosbash, M. (2004). Coupled oscillators control morning and evening locomotor behaviour of Drosophila. Nature 431, 862-868.

5.

Dubruille, R., and Emery, P. (2008). A Plastic Clock: How Circadian Rhythms Respond to Environmental Cues in Drosophila. Molecular neurobiology 38, 129-145.

6.

Koh, K., Zheng, X., and Sehgal, A. (2006). JETLAG resets the Drosophila circadian clock by promoting light-induced degradation of TIMELESS. Science 312, 1809-1812.

7.

Calgaro, S., Boube, M., Cribbs, D.L., and Bourbon, H.M. (2002). The Drosophila gene taranis encodes a novel trithorax group member potentially linked to the cell cycle regulatory apparatus. Genetics 160, 547-560.

8.

Hsu, S.I., Yang, C.M., Sim, K.G., Hentschel, D.M., O'Leary, E., and Bonventre, J.V. (2001). TRIP-Br: a novel family of PHD zinc finger- and

78

bromodomain-interacting proteins that regulate the transcriptional activity of E2F-1/DP-1. The EMBO journal 20, 2273-2285. 9.

Afonso, D.J., Liu, D., Machado, D.R., Pan, H., Jepson, J.E., Rogulja, D., and Koh, K. (2015). TARANIS Functions with Cyclin A and Cdk1 in a Novel Arousal Center to Control Sleep in Drosophila. Current biology : CB 25, 17171726.

10.

Emery, P., So, W.V., Kaneko, M., Hall, J.C., and Rosbash, M. (1998). CRY, a Drosophila clock and light-regulated cryptochrome, is a major contributor to circadian rhythm resetting and photosensitivity. Cell 95, 669-679.

11.

Yao, Z., and Shafer, O.T. (2014). The Drosophila circadian clock is a variably coupled network of multiple peptidergic units. Science 343, 1516-1520.

12.

Stoleru, D., Peng, Y., Nawathean, P., and Rosbash, M. (2005). A resetting signal between Drosophila pacemakers synchronizes morning and evening activity. Nature 438, 238-242.

13.

Shafer, O.T., and Yao, Z. (2014). Pigment-Dispersing Factor Signaling and Circadian Rhythms in Insect Locomotor Activity. Current opinion in insect science 1, 73-80.

14.

Taghert, P.H., and Nitabach, M.N. (2012). Peptide neuromodulation in invertebrate model systems. Neuron 76, 82-97.

15.

Kula, E., Levitan, E.S., Pyza, E., and Rosbash, M. (2006). PDF cycling in the dorsal protocerebrum of the Drosophila brain is not necessary for circadian clock function. Journal of biological rhythms 21, 104-117.

16.

Park, J.H., and Hall, J.C. (1998). Isolation and chronobiological analysis of a neuropeptide pigment-dispersing factor gene in Drosophila melanogaster. Journal of biological rhythms 13, 219-228.

79

17.

Abruzzi, K.C., Rodriguez, J., Menet, J.S., Desrochers, J., Zadina, A., Luo, W., Tkachev, S., and Rosbash, M. (2011). Drosophila CLOCK target gene characterization: implications for circadian tissue-specific gene expression. Genes & development 25, 2374-2386.

18.

Kim, E.Y., Bae, K., Ng, F.S., Glossop, N.R.J., Hardin, P.E., and Edery, I. (2002). Drosophila CLOCK Protein Is under Posttranscriptional Control and Influences Light-Induced Activity. Neuron 34, 69-81.

19.

Lu, B., Liu, W., Guo, F., and Guo, A. (2008). Circadian modulation of lightinduced locomotion responses in Drosophila melanogaster. Genes, Brain and Behavior 7, 730-739.

20.

Kumar, S., Chen, D., and Sehgal, A. (2012). Dopamine acts through Cryptochrome to promote acute arousal in Drosophila. Genes & development 26, 1224-1234.

21.

Wheeler, D.A., Hamblen-Coyle, M.J., Dushay, M.S., and Hall, J.C. (1993). Behavior in Light-Dark Cycles of Drosophila Mutants That Are Arrhythmic, Blind, or Both. Journal of biological rhythms 8, 67-94.

22.

Fernandez, M.P., Berni, J., and Ceriani, M.F. (2008). Circadian remodeling of neuronal circuits involved in rhythmic behavior. PLoS Biol 6, e69.

23.

Sivachenko, A., Li, Y., Abruzzi, K.C., and Rosbash, M. (2013). The transcription factor Mef2 links the Drosophila core clock to Fas2, neuronal morphology, and circadian behavior. Neuron 79, 281-292.

24.

Petsakou, A., Sapsis, T.P., and Blau, J. (2015). Circadian Rhythms in Rho1 Activity Regulate Neuronal Plasticity and Network Hierarchy. Cell 162, 823835.

80

25.

Zang, Z.J., Gunaratnam, L., Cheong, J.K., Lai, L.Y., Hsiao, L.-L., O'Leary, E., Sun, X., Salto-Tellez, M., Bonventre, J.V., and Hsu, S.I.H. (2009). Identification of PP2A as a novel interactor and regulator of TRIP-Br1. Cellular signalling 21, 34-42.

26.

Sathyanarayanan, S., Zheng, X., Xiao, R., and Sehgal, A. (2004). Posttranslational regulation of Drosophila PERIOD protein by protein phosphatase 2A. Cell 116, 603-615.

27.

Viquez, N.M., Fuger, P., Valakh, V., Daniels, R.W., Rasse, T.M., and DiAntonio, A. (2009). PP2A and GSK-3beta act antagonistically to regulate active zone development. The Journal of neuroscience : the official journal of the Society for Neuroscience 29, 11484-11494.

28.

Yoshii, T., Wulbeck, C., Sehadova, H., Veleri, S., Bichler, D., Stanewsky, R., and Helfrich-Forster, C. (2009). The neuropeptide pigment-dispersing factor adjusts period and phase of Drosophila's clock. The Journal of neuroscience : the official journal of the Society for Neuroscience 29, 2597-2610.

29.

Cusumano, P., Klarsfeld, A., Chelot, E., Picot, M., Richier, B., and Rouyer, F. (2009). PDF-modulated visual inputs and cryptochrome define diurnal behavior in Drosophila. Nature neuroscience 12, 1431-1437.

30.

Kim, E.Y., Bae, K., Ng, F.S., Glossop, N.R., Hardin, P.E., and Edery, I. (2002). Drosophila CLOCK protein is under posttranscriptional control and influences light-induced activity. Neuron 34, 69-81.

31.

Kumar, S., Chen, D., and Sehgal, A. (2012). Dopamine acts through Cryptochrome to promote acute arousal in Drosophila. Genes & development 26, 1224-1234.

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32.

Allada, R., White, N.E., So, W.V., Hall, J.C., and Rosbash, M. (1998). A Mutant Drosophila Homolog of Mammalian Clock Disrupts Circadian Rhythms and Transcription of period and timeless. Cell 93, 791-804.

33.

Jepson, J.E., Shahidullah, M., Lamaze, A., Peterson, D., Pan, H., and Koh, K. (2012). dyschronic, a Drosophila homolog of a deaf-blindness gene, regulates circadian output and Slowpoke channels. PLoS Genet 8, e1002671.

82

A 8

16

0

8

elav > +

elav > tara RNAi

1122rhyM017C14.txt 16

0 0

8

16

0

1122rhyM022C09.txt

8

16

0 0

1

1

2

2

2

3

3

3

4

4

4

5

5

5

6

6

6

7

7

7

8

8

8

DD

1

C

1207rhyM009C06.txt 0

8

16

+/+

1207rhyM010C02.txt

0

8

0 0

16

1

2

2

3

3

4

4

5

5

DD

1

e/s

6

1129rhyM069C22.txt 70

E

8

16

tara 0

8

6

1 8

2

2

3

3

4

4

5

5

6

6

Genotype 8 Knockdown + > tara RNAi elav > + elav > tara RNAi Rescue +/+ Ub-tara tarae/s Ub-tara; tarae

N

7

8

16

0

8

16

8

0

16

16

0

8

16

0

8

16

0

0

160 120 80 ***

40 0

i Ai > + NA N R v R a ela ra r ta ta > > + av el

D

e

Ub-tara; tara

1207rhyM008C09.txt

0 70

16

81

7

Ub-tara

8

Rhythm strength

+ > tara RNAi

1129rhyM029C06.txt 0

Rhythm strength

B

160 120

***

80 40 0

l e /s ro ara ae ra t n -t r a co Ub ta a; t r ta Ub

%R

% WR

% AR

45 53 73

100 88.7 26

0 3.8 21.9

0 7.5 52.1

24.4 ± 0.07 24.3 ± 0.17 23.9 ± 0.18

117.1 ± 4.65 98.3 ± 5.14 30.7 ± 4.73

48 48 26 49

95.8 89.6 50.0 85.7

4,2 8.3 23.1 10.2

0 2.1 26.9 4.1

24.1 ± 0.08 24.2 ± 0.12 24.1 ± 0.15 25.1 ± 0.08

116.8 ± 4.67 96.4 ± 5.10 44.8 ± 8.50 99.4 ± 5.45

8

tau (h) ± SEM

Power ± SEM

N: number of flies; R: rhythmic; WR: weakly rhythmic; AR: arrhythmic tau: free-running period; Power: measure rhythm strength;

Figure 1

83

8

16

0

8

1005rhyM028C24.txt 16

0 0

8

tim > + 16

0

1005rhyM026C23.txt

8

16

0 0

1

1

1

2

2

2

3

3

3

4

4

4

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5

5

6

6

6

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7

7

8

8

8

DD

tim > tara RNAi

Rhythm strength

+ > tara RNAi

1005rhyM027C18.txt 0

B

8

16

0

8

16

0

D + > tara RNAi

0421rhyM030C29.txt 8

Scale: 0.0

16

0

8

16

6.3 0

Pdf > +

Pdf > tara RNAi

DD

1

2

3

4

80

***

40 0

160 120 80 40 0

i Ai > + NA N R df R P ra a r ta ta > > f + Pd

5

6

7

8

E

120

i + Ai A > N RN tim a R r ra ta ta > > + it m

C 0

160

Rhythm strength

A

Genotype

N

%R

% WR

% AR

tau(h) ± SEM

Power ± SEM

+ > tara RNAi tim > + tim > tara RNAi

46 47 43

97.8 100 72.1

2.2 0 14

0 0 14

23.8 ± 0.06 24.3 ± 0.05 24.2 ± 0.08

141.3 ± 5.12 135.6 ± 3.23 69.3 ± 5.99

+ > tara RNAi Pdf > + Pdf > tara RNAi

31 31 47

100 100 100

0 0 0

0 0 0

23.6 ± 0.05 24.3 ± 0.06 24.0 ± 0.05

144.9 ± 4.90 137.5 ± 2.98 131.6 ± 3.04

N: number of flies; R: rhythmic; WR: weakly rhythmic; AR: arrhythmic tau: free-running period; Power: measure rhythm strength;

Figure 2 84

A

Pdf > +

Pdf > tara FL

DD

+ > tara FL

D

C

160 120 80 40 0

FL > + FL ra df ara a t P t > f> + d P

Genotype

N

+ > tara FL Pdf > + Pdf > tara FL

32 32 18

Period length (h)

Rhythm strength

B

27 26 25 24 23

L

FL > + a F a f r r ta Pd > ta > f + Pd

%R 96.9 96.9 77.8

% WR 3.1 3.1 16.6

% AR

tau(h) ± SEM

Power ± SEM

0 0 5.5

24.7 ± 0.12 24.7 ± 0.10 26.1 ± 0.28

118.0 ± 6.03 102.4 ± 6.15 89.7 ± 8.83

N: number of flies; R: rhythmic; WR: weakly rhythmic; AR: arrhythmic tau: free-running period; Power: measure rhythm strength;

Figure 3

85

control

CT3 PER PDF

DD3: sLNvs CT15 CT9

1/s

tara

C

DD3: DN1s

control

PER PDF

1/s

tara

Figure 4

86

CT21

B PER signal (a.u.)

A

1.2 0.8 0.4 0

***

control 1/s ns tara

ns

3 9 15 21 Circadian time

CT3 PER PDF

+ > tara FL

Pdf > +

DD3: sLNvs CT9 CT15

B CT21

PER signal (a.u.)

A

+ > tara FL Pdf > + Pdf > tara FL

1.2 0.8 0.4 0

*** 3

9 15 21 Circadian time

Pdf > tara FL

Figure 5

87

A PDF

DD3: sLNv dorsal projection CT9 CT15

CT3

CT21

control

1/s

tara

IF intensity

IF intensity (a.u.)

1.2

control 1/s tara

0.8 0.4 0

Activity levels/ 30 min

D 100

50

0

Figure 6

88

*** 3

***

9 15 21 Circadian time control

C Pdf mRNA (a.u.)

B

1.2

**

0.8 0.4 0

l

n co

tro

ZT15

s

1/

ra

ta

1/s

tara

A

Pdf > taraFL

Pdf > +

Pdf > tara RNAi

C 100

100 0 f Pd

>

+

f>

Pd

ta

r

L aF

f>

+

Pd ra ta > f Pd

RN

Ai

80 60

*

200

Pdf > mCD8::GFP

*

Pdf > mCD8::GFP

Branching points

300

ZT3

Pdf > +

***

Projection length (µm)

B

Pdf > mCD8::GFP

40 20 0 f>

Pd

+

Pd

f>

L

ta

F ra

f>

+

Pd ra ta > f Pd

i

A RN

Figure 7

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FIGURE LEGENDS

Figure 1. Pan-neuronal tara knockdown degrades while ubiquitous rescue restores rhythm strength. (A) tara pan-neuronal knockdown markedly reduces rhythm strength. Representative double-plotted actograms of single background control (+ > tara RNAi and elav > +) and knockdown (elav > tara RNAi) female flies are shown. Only 2 male flies elav > tara RNAi survived for six days in free-running conditions precluding the analysis of their rhythmic behavior. Gray and black bars above the actogram indicate subjective day and night, respectively. Six 6 days in free-running conditions (constant darkness (DD) and temperature condition) are shown. (B) Rhythm strength of control flies (+ > tara RNAi and elav > +) and knockdown flies (elav > tara RNAi). Mean ± SEM is shown. (C) Representative double-plotted actograms for female flies of the indicated genotypes. (D) Rhythm strength for the same genotypes shown in (C). Mean ± SEM is shown. (E) Rest:activity locomotor rhythms for same flies shown in (B) and (D). . ***p < 0.001, one-way ANOVA followed by post hoc tests: Dunnett (B) and Tukey (D). tau indicates period length for rhythmic flies in each genotype.

Figure 2. tara knockdown specifically in the clock cell network degrades locomotor rhythms. (A) tara knockdown in clock cells markedly reduces rhythm strength. Representative double-plotted actograms of background control (+ > tara RNAi and tim > +) and knockdown (tim > tara RNAi) male flies are shown. Gray and black bars above the actogram indicate subjective day and night, respectively. Six 6 days in free-running conditions (constant darkness (DD) and temperature condition) are

90

shown. (B) Rhythm strength of control flies (+ > tara RNAi and tim > +) and knockdown flies tim > tara RNAi. Mean ± SEM is shown. (C) tara knockdown in PDF+ cells does not impact rhythm strength. Representative double-plotted actograms of background control (+ > tara RNAi and Pdf > +) and knockdown (Pdf > tara RNAi) male flies are shown. (D) Rhythm strength of control flies (+ > tara RNAi and Pdf > +) and knockdown flies Pdf > tara RNAi. Mean ± SEM is shown. (E) Rest:activity locomotor rhythms for the same flies shown in (B) and (D). ***p < 0.001 one-way ANOVA followed by Dunnett post hoc test (B).

Figure 3. tara overexpression in PDF+ cells increases period length of locomotor activity without significantly degrading rhythm strength. (A) Representative double-plotted actograms of background control (+ > tara FL and Pdf > +) and overexpression (Pdf > tara FL) female flies are shown. Gray and black bars above the actogram indicate subjective day and night, respectively. Six 6 days in free-running conditions (constant darkness (DD) and temperature condition) are shown. (B) Rhythm strength for the indicated genotypes. (C) Period length for the indicated genotypes. Mean ± SEM is shown. (D) Rest:activity locomotor rhythms for same flies shown in (B).

Figure 4. In tara mutants PER cycling in sLNvs is faster. (A) PERIOD and PDF costaining in sLNvs for control and tara1/s at the circadian times shown after three days in free running conditions. Scale bar is 10µm. (B) PERIOD signal quantification in sLNvs for control (n=16 to18) and tara1/s (n=11 to 18) male brains. Mean ± SEM is shown. (C) PERIOD and PDF co-staining in DN1 neurons for control and tara1/s at the indicated

91

circadian times shown, after three days in free running conditions. Scale bar is 20 µm. ***p < 0.001, ns: not significant, one-way ANOVA followed by Tukey post hoc test (B).

Figure 5. TARA overexpression in PDF+ neurons negatively regulates the pace of molecular clock. (A) PERIOD and PDF co-staining in sLNvs for controls (+ > tara FL and Pdf > +) and overexpression (Pdf > tara FL) male flies at the indicated circadian times after three days in free running conditions. Scale bar is 10µm. (B) PERIOD signal quantification in sLNvs for + > tara FL (n=8 to 11), Pdf > + (n=10 to 16), and Pdf > tara FL (n=10-12). Mean ± SEM is shown. ***p < 0.001, one-way ANOVA followed by Tukey post hoc test (B).

Figure 6. tara regulates Pdf both transcriptional and post-transcriptionally. (A) Representative sLNv dorsal projection at the indicated circadian times are shown after three

days

in

free

running

conditions

for

control

and

tara1/s

male

flies.

Immunofluorescence (IF) intensity is artificially represented by a color spectrum where black represents the lowest intensity and white represents the maximum intensity. Scale bar is 20µm. (B) PDF signal quantification in the sLNv dorsal projection for control (n=16 to 18) and for tara1/s (n=15 to 17). Mean ± SEM is shown. (C) Pdf mRNA levels in control and tara1/s flies. Average of three independent sets is shown. Mean ± SEM is shown. (D) Morning and evening anticipation activities for control and tara1/s flies in LD conditions. ***p < 0.001, one-way ANOVA followed by Tukey post hoc test (B). **p < 0.01, t test analysis (C).

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Figure 7. tara modulates sLNv dorsal projection structural morphology. (A) Representative sLNv dorsal projection for the indicated genotypes are shown. sLNv dorsal projection were reconstructed from confocal microscope files using the neurostudio software. To label the entire sLNv dorsal projection we made use of membrane targeted GFP construct (UAS-mCD8::GFP). (B) Quantification of sLNv dorsal projection length quantification for the same genotypes shown in (A). Quantification was automatic starting on the first branching point to the tip of the projection. Pdf > + (n=10), Pdf > tara FL (n=10), Pdf > dicer2 (n=10), and Pdf > dicer2; tara RNAi (n=6). (C) Number of branching points for the same genotypes shown in (A). Mean ± SEM is shown. *p < 0.05, and ***p < 0.001, t test analysis.

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CT3 GFP

GFP PDF

DD3: sLNvs

CT9

CT15

CT21

B TARA::GFP (a.u.)

A

1.6 1.2

ns

0.8 0.4 0

3 9 15 21 Circadian time

Figure S1. TARA protein levels do not cycle in circadian pacemaker neurons. (A) Immunostaining of TARA::GFP in male fly brains on the 3rd day in DD. We used transgenic flies that carry an artificial exon encoding GFP inserted into an intron of tara in the genome and therefore are expected to produce endogenous levels of TARA protein fused to GFP. Brains were dissected at indicated circadian times (CT) and stained for GFP (green) and PDF (red), which was used to identify small ventral lateral neurons (sLNvs), the pacemaker neurons in DD. Scale bar 10µm. (B) Quantification of TARA::GFP signal in sLNvs. Data from 11-16 brain hemispheres are presented. Mean ± SEM is shown. ns: not significant, two-way ANOVA followed by Tukey post hoc test (B).

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Chapter 3 _______________________________________________________________ Discussion, Conclusions and Future Perspectives

I.

tara is a novel regulator of sleep

Our results demonstrate that tara has an important role in sleep regulation. tara mutants display a dramatic decrease in daily sleep during the day and the night. They also have an insomnia-like phenotype with increased latency in sleep initiation. These defects can be rescued with the introduction of tara cDNA in the tara mutant background. We also found that TARA is expressed in neurons but excluded from glial cells. Importantly, the role of tara in sleep regulation is, in part, adult specific. Sleep is controlled mainly by two mechanisms: a circadian mechanism that times sleep to an ecologically relevant part of the day, and a homeostatic mechanism that ensures that an adequate amount of sleep is maintained [1]. Most of the stronger tara mutants are arrhythmic, yet sleep levels are decreased in constant light and constant dark conditions which demonstrates that tara regulates sleep independently of the circadian mechanism and of the light input pathways. These observations leave the homeostatic mechanism as the probable place where tara function is required to control sleep amount. Indeed our preliminary data suggest that TARA impacts the homeostatic response to sleep deprivation. When deprived of sleep for 4 hours toward the end of the night, tara mutants exhibited significantly reduced rebound sleep compared to control flies. However, after 12 hours of sleep deprivation, the rebound sleep of tara mutants was similar to that of the controls (data not shown). These data demonstrate that tara mutants exhibit a dose-dependent homeostatic response to sleep deprivation. Additional studies are required to determine the mechanism by which TARA controls sleep homeostasis.

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II.

tara, CycA, and Cdk1 form a new sleep regulatory pathway

TARA has a CycA binding homology domain and CycA was shown to regulate sleep [2]. These led us to hypothesize tara and CycA regulate sleep through the same genetic pathway. Our genetic and physical interaction studies demonstrate that tara and CycA indeed interact with each other to regulate sleep. This is important because we did not simply find a new sleep regulatory molecule, but also demonstrated how this new molecule is integrated into a larger gene network consisting of cell cycle genes [2]. TARA was previously described as a transcriptional co-regulator [3]. This previous work and our finding that TARA::GFP fusion protein is expressed in neuronal nuclei are consistent with a role for tara in transcriptional control. However, TARA physically binds CycA and regulates its levels at the posttranscriptional level. As discussed in more detail below, these observations suggest that TARA regulates sleep through a novel non-transcriptional mechanism independent from those controlling cell cycle progression. Future experiments aimed at separating the transcriptional from the non-transcriptional roles may help determine their relative importance for sleep regulation. Through bioinformatics analysis, Calgaro et. al showed that TARA has four conserved domains [3], one of which being the C-terminal domain.

In the

mammalian homologs of TARA, the C-terminal domain appears to be required for their transcriptional role [4]. We hypothesize that TARA controls sleep through a non-transcriptional mechanism which will be clarified by studying whether the C-terminal domain of TARA is required for transcriptional regulation but dispensable for sleep regulation.

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Our data suggest that CycA regulates sleep through its modulatory action over Cdk1 activity. Cdk1 is regulated through inhibitory phosphorylation of its T14 and Y15 residues, which is dependent on the cellular context [5]. Because overexpression of wild-type Cdk1 did not affect sleep, presumably due to tight regulatory mechanisms, we made use of the Cdk1 (T14A,Y15F) mutant, termed Cdk1-AF, that cannot be inhibited [5] to show that increased activity of Cdk1 suppresses sleep. Our model is that CycA, and possibly other cyclins, suppresses Cdk1 activity within the CycA+ cells, reducing its activity and therefore increasing sleep. Other scenarios are possible, but a direct relationship between CycA and Cdk1 is likely, given that CycA and Cdk1 are known to physically interact [6]. Interestingly, both CycA and Cdk1 localize to synaptic regions, which suggest a direct modulatory role for CycA and Cdk1 over synaptic proteins. Identification of the substrates of the Cdk1 kinase activity relevant for sleep regulation is an important next step we intend to pursue in future experiments. Recent work in Drosophila has demonstrated that knockdown of Cdk1 and other Cyclin/Cdk family genes significantly rescues seizure duration in both bas1 (bang sensitive1) and bss1 (bang sensless1) mutants [7]. This suggests that Cdk1 may modulate ion channel activity and membrane excitability. Several lines of evidence show that ion channels have a dramatic influence over sleep. Shaker, a fast delayed rectifier potassium channel [8], hyperkinetic, the Shaker cytoplasmic beta subunit [9], ether-à-go-go, slow delayed rectifier potassium channel [10], redeye, nicotinic Acetylcholine Receptor α4 [11], and Rdl, a GABAA receptor gene [12] are all implicated in sleep and may be potential targets for Cdk1. Given that increased excitability of PL neurons suppresses

99

sleep, the net effect of Cdk1 activity over these channels may be an overall increase in neuronal excitability. Thus, these channels may serve as plausible phosphorylation targets of Cdk1. The relationship between sleep and epilepsy is well established [13]. However, how sleep and epilepsy are interconnected at the molecular level is less clear. Interestingly, cell cycle genes are implicated in both sleep and epilepsy. Therefore, addressing how the cell cycle regulators tara and Cdk1 regulate sleep may contribute to our understanding of how epileptogenic processes develop, highlighting how the finding of a basic molecular mechanism regulating one behavior contributes to the understanding of other processes.

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III.

A novel arousal circuit modulated by tara, CycA, and Cdk1

Previous work showed that CycA protein is expressed in a small number of neurons [2]. In order to manipulate the CycA expressing cells we made use of the PL (pars lateralis)-Gal4, which labels just the dorsal cluster of CycA expressing cells. Increasing the activity of these neurons led to a strong sleep suppression phenotype, suggesting that they serve as an arousal center. Importantly, we demonstrated their adult function by showing that adult-specific and reversible activation of these neurons suppresses sleep. Given that constitutive expression of Cdk1-AF phenocopies PL neurons activation, we envision a model where CycA regulates the activity of PL neurons through Cdk1. In CycA mutants, lower levels of CycA lead to higher Cdk1 activity, which in itself increases the excitability of PL neurons and, therefore, promotes wakefulness. Whether or not Cdk1 leads to an overall increase in the excitability of PL neurons is of interest to address in the future. Different groups of neuronal populations are implicated in the regulation of sleep. These include the mushroom body [14-16], the fan shaped body [17], the pars intercerebralis [18], the dorsal paired medial (DPM) neurons [19], the PPL1 cluster of TYROSINE HYDROXYLASE positive neurons [20], and the large ventral lateral clock neurons (lLNvs) [21, 22]. For future directions, it will be interesting to determine whether or not PL neurons form synapses with previously identified sleep/arousal centers. In order to address that, we can make use of GFP reconstitution across synaptic partners (GRASP). If PL neurons form synapses with other known sleep relevant neuronal population, it will be of further interest to examine how changes in the neuronal activity of one

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group of neurons modify the activity of the other group using genetically encoded Ca2+ indicators [23].

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IV.

tara may control sleep by regulating synaptic structure

During sleep animals cannot engage in productive behaviors as foraging or mating and are at a higher vulnerability to predation. This apparent weakness of sleep contrasts with its near universality across different organisms, which suggests that the benefits of sleep outweigh the hazards [24]. Sleep may have more than one function; it may restore energy and supplies in the brain, rescale synapses, and consolidate memories [25]. Increasing evidence suggests that sleep is important for neural plasticity [26, 27]. In rodents, markers of synaptic strength are higher after wake and lower after sleep [28, 29]. Evidence from Drosophila has also shown that the volume of synapses in the sLNv dorsal projection is increased after wakefulness and sleep deprivation [26], establishing the sLNv dorsal projection as a useful neuronal target for the study of sleep dependent plastic changes. Our results showing that both CycA and Cdk1 localize to the synaptic terminals suggest that they play a role in synaptic function. TARA may play a role in synaptic morphology through its interaction with CycA and Cdk1. Our data demonstrating that tara knockdown in PDF+ neurons dramatically affects dorsal projection morphology is proof of principle that TARA affects synaptic structure. The molecular mechanisms underlying the role of TARA in synaptic structure and function may be a fruitful area of future research.

103

V.

tara negatively regulates the intrinsic speed of the molecular clock

The molecular clock synchronizes organismal physiology to external environmental conditions at a pace of ~24 hours. Different genes and circuits are responsible for maintaining the speed of the molecular clock even in the absence of external cues. Our initial analysis of circadian locomotor activity demonstrated that most of the stronger tara mutant flies were arrhythmic, which led us to ask whether the molecular clock was still cycling. Previously we reported that PER still cycles robustly in the stronger tara mutants in LD conditions [30]. Our new data revealed that PER protein cycles at a faster pace in tara mutants than in control flies in DD conditions, suggesting that TARA normally decreases the speed of the molecular clock. We did not detect a short-period locomotor rhythm in tara mutants, but this could be because more severely affected mutants become arrhythmic, making it difficult to determine their period length. Our finding that TARA overexpression in PDF+ neurons slows down PER cycling and increases the period length of locomotor rhythm further points to a negative effect of TARA in the intrinsic speed of the molecular clock. Multiple interlocking transcriptional feedback loops that regulate the pace of the molecular clock were identified in Drosophila. In addition to the main loop involving PER/TIM, CLK/CYC directly activates the transcription of their activator PAR domain protein 1 (Pdp1) as well as the repressors vrille (vri) and clockwork

orange

(cwo)

[31].

Previous

chromatin

immunoprecipitation

experiments suggest that tara is a direct transcriptional target of CLK [32]. While we did not observe cycling of tara mRNA levels in whole head extracts, tara

104

function may cycle in clock neurons. Given the known role of TARA in transcriptional regulation, it will be interesting to investigate whether TARA interacts with previously described feedback loops or functions in a novel transcriptional feedback loop. Most strong tara mutants are arrhythmic but exhibit robust molecular cycling of PER, which suggests that tara may play other roles downstream of the molecular clock. In contrast to the core clock mechanism, only a limited number of molecules were shown to act downstream of the core molecular clock, and those are: Pigment dispersing factor (Pdf) [33, 34], Pdf receptor [35-37], neurofibromatosis1 [38], slowpoke [39], narrow abdomen [40], ebony [41], miR279 [42], dyschronic [43], and diuretic hormone 44 [44]. Starting from such a selected group of genes will simplify future studies investigating genetic interactions between tara and genes that are pivotal to the output arm of the circadian mechanism.

105

VI.

tara regulates PDF levels

The circadian locomotor rhythm of Pdf null mutants dampens in free-running conditions and their morning anticipation is absent while the evening anticipation is advanced [34]. tara mutants exhibit low levels of PDF in the sLNv dorsal projection, yet tara mutants have close to normal morning and evening anticipations in LD conditions. This suggests that low levels of PDF are sufficient to drive morning and evening anticipations in these mutants. Examination of tara mutants in long photoperiod conditions, where Pdf null mutants are unable to delay the phase of the evening peak [45], may provide a more sensitive assay for the functional consequences of low PDF levels. Robust circadian locomotor activity rhythms rely on the entire clock cell network. However, the sLNvs set the pace of the other clock neurons in constant darkness as long as their pace is not more than 2.5 hours apart from the pace of the sLNvs [46]. This synchronizing effect is largely mediated by PDF. PDF is expressed in the sLNvs and lLNvs, which represent only 16-18 cells out of ~150 clock neurons. The sLNv dorsal projection is in close proximity to the dorsal groups of clock cells and it was shown that more than 60% of the clock cells respond to PDF [47]. Daily rhythms in the morphology of the sLNv dorsal projection were described [48], which include not only daily cycles in fasciculation and defasciculation [49], but

also gains and losses in axonal material [50].

However, the relevance of these daily structural changes remains unclear [51]. Our results showing that flies with knockdown of TARA in sLNvs are rhythmic despite exhibiting altered morphology of the sLNv dorsal projection suggest that

106

the morphological changes are not the primary cause of the circadian rhythm phenotypes in tara mutants. Still, low levels of PDF in these neurons likely contribute to the desynchronization of the clock cell network and to the reduced rhythm strength seen in tara mutants.

107

VII.

tara regulates nocturnal activity and neuronal morphology

As humans, flies are diurnal organisms. Interestingly, tara mutants display increased nighttime activity suggesting that tara is important for the regulation of the diurnal pattern of activity. Clk mutants also have an increased nocturnal activity phenotype [52, 53], and physically bind tara genomic locus [32], raising the possibility that Clk and tara work in the same genetic pathway to regulate the diurnal pattern of behavior of flies. In Clkjrk mutants, CRYPTOCHROME (CRY) and dopamine levels are elevated [54], Thus, we will next test whether blocking dopamine signaling and CRY activity in tara mutants restore their diurnal activity, and whether tara and Clk genetically interact to regulate nighttime activity. Another interesting avenue for future research derives from a previous finding that TRIP-Br1, one of the tara mammalian homologs, physically interacts with and is regulated by PROTEIN PHOSPHATASE 2A (PP2A) [55]. In mammalian cells, overexpression of PP2A decreased the phosphorylated form of TRIP-Br1 and increased total TRIP-Br1 protein levels, suggesting that PP2A is a positive regulator of TRIP-Br1 [55]. Interestingly, PP2A plays a role in the molecular clock by directly dephosphorylating PER and controlling its stability and cycling [56]. PP2A is an abundant heterotrimeric enzyme composed of a highly conserved catalytic subunit, a variable regulatory subunit, and a structural subunit. In Drosophila, there are four genes encoding PP2A regulatory subunits, two of which, twins and widerborst, are under circadian control [56].

108

In addition to its role in the molecular clock, PP2A regulates active zone development and synaptic terminal morphology at the Drosophila NMJ [57]. This role of PP2A in synaptic morphology is particularly interesting in light of our results suggesting that tara regulates the morphology of sLNv dorsal projection. It will be interesting to examine whether PP2A and TARA interact to regulate synaptic morphology and locomotor rhythms.

In conclusion, we have identified two new sleep regulatory molecules that, together with other cell cycle genes, may become important targets for the treatment of sleep disorders. Importantly, tara, CycA and Cdk1 have mammalian homologs that may regulate sleep as well. Beyond that, this work constitutes a strong foundation for future lines of research, including the understanding of diurnal versus nocturnal behaviors and aspects of the structure and function of synapses.

109

VIII.

References

1.

Borbely, A.A., and Achermann, P. (1999). Sleep homeostasis and models of sleep regulation. Journal of biological rhythms 14, 557-568.

2.

Rogulja, D., and Young, M.W. (2012). Control of sleep by cyclin A and its regulator. Science 335, 1617-1621.

3.

Calgaro, S., Boube, M., Cribbs, D.L., and Bourbon, H.M. (2002). The Drosophila gene taranis encodes a novel trithorax group member potentially linked to the cell cycle regulatory apparatus. Genetics 160, 547-560.

4.

Hsu, S.I., Yang, C.M., Sim, K.G., Hentschel, D.M., O'Leary, E., and Bonventre, J.V. (2001). TRIP-Br: a novel family of PHD zinc finger- and bromodomain-interacting proteins that regulate the transcriptional activity of E2F-1/DP-1. The EMBO journal 20, 2273-2285.

5.

Ayeni, J.O., Varadarajan, R., Mukherjee, O., Stuart, D.T., Sprenger, F., Srayko, M., and Campbell, S.D. (2014). Dual phosphorylation of cdk1 coordinates cell proliferation with key developmental processes in Drosophila. Genetics 196, 197-210.

6.

Meyer, C.A., Jacobs, H.W., Datar, S.A., Du, W., Edgar, B.A., and Lehner, C.F. (2000). Drosophila Cdk4 is required for normal growth and is dispensable for cell cycle progression. The EMBO journal 19, 4533-4542.

7.

Lin, W.-H., He, M., and Baines, R.A. (2015). Seizure suppression through manipulating splicing of a voltage-gated sodium channel. Brain 138, 891-901.

110

8.

Cirelli, C., Bushey, D., Hill, S., Huber, R., Kreber, R., Ganetzky, B., and Tononi, G. (2005). Reduced sleep in Drosophila Shaker mutants. Nature 434, 1087-1092.

9.

Bushey, D., Huber, R., Tononi, G., and Cirelli, C. (2007). Drosophila Hyperkinetic mutants have reduced sleep and impaired memory. The Journal of neuroscience : the official journal of the Society for Neuroscience 27, 5384-5393.

10.

Rihel, J., Prober, D.A., Arvanites, A., Lam, K., Zimmerman, S., Jang, S., Haggarty, S.J., Kokel, D., Rubin, L.L., Peterson, R.T., et al. (2010). Zebrafish behavioral profiling links drugs to biological targets and rest/wake regulation. Science 327, 348-351.

11.

Shi, M., Yue, Z., Kuryatov, A., Lindstrom, J.M., and Sehgal, A. (2014). Identification of Redeye, a new sleep-regulating protein whose expression is modulated by sleep amount, Volume 3.

12.

Chung, B.Y., Kilman, V.L., Keath, J.R., Pitman, J.L., and Allada, R. (2009). The GABA(A) receptor RDL acts in peptidergic PDF neurons to promote sleep in Drosophila. Current biology : CB 19, 386-390.

13.

Derry, C.P., and Duncan, S. (2013). Sleep and epilepsy. Epilepsy & Behavior 26, 394-404.

14.

Joiner, W.J., Crocker, A., White, B.H., and Sehgal, A. (2006). Sleep in Drosophila is regulated by adult mushroom bodies. Nature 441, 757-760.

15.

Pitman, J.L., McGill, J.J., Keegan, K.P., and Allada, R. (2006). A dynamic role for the mushroom bodies in promoting sleep in Drosophila. Nature 441, 753-756.

111

16.

Sitaraman, D., Aso, Y., Jin, X., Chen, N., Felix, M., Rubin, G.M., and Nitabach, M.N. (2015). Propagation of Homeostatic Sleep Signals by Segregated Synaptic Microcircuits of the Drosophila Mushroom Body. Current biology : CB.

17.

Donlea, J.M., Thimgan, M.S., Suzuki, Y., Gottschalk, L., and Shaw, P.J. (2011). Inducing sleep by remote control facilitates memory consolidation in Drosophila. Science 332, 1571-1576.

18.

Crocker, A., Shahidullah, M., Levitan, I.B., and Sehgal, A. (2010). Identification of a neural circuit that underlies the effects of octopamine on sleep:wake behavior. Neuron 65, 670-681.

19.

Haynes, P.R., Christmann, B.L., and Griffith, L.C. (2015). A single pair of neurons links sleep to memory consolidation in Drosophila melanogaster. Elife 4.

20.

Liu, Q., Liu, S., Kodama, L., Driscoll, M.R., and Wu, M.N. (2012). Two dopaminergic neurons signal to the dorsal fan-shaped body to promote wakefulness in Drosophila. Current biology : CB 22, 2114-2123.

21.

Sheeba, V., Fogle, K.J., Kaneko, M., Rashid, S., Chou, Y.T., Sharma, V.K., and Holmes, T.C. (2008). Large ventral lateral neurons modulate arousal and sleep in Drosophila. Current biology : CB 18, 1537-1545.

22.

Parisky, K.M., Agosto, J., Pulver, S.R., Shang, Y., Kuklin, E., Hodge, J.J., Kang, K., Liu, X., Garrity, P.A., Rosbash, M., et al. (2008). PDF cells are a GABA-responsive wake-promoting component of the Drosophila sleep circuit. Neuron 60, 672-682.

112

23.

Riemensperger, T., Pech, U., Dipt, S., and Fiala, A. (2012). Optical calcium imaging in the nervous system of Drosophila melanogaster. Biochimica et biophysica acta 1820, 1169-1178.

24.

Allada, R., and Siegel, J.M. (2008). Unearthing the phylogenetic roots of sleep. Current biology : CB 18, R670-R679.

25.

Tononi, G., and Cirelli, C. (2014). Sleep and the price of plasticity: from synaptic and cellular homeostasis to memory consolidation and integration. Neuron 81, 12-34.

26.

Bushey, D., Tononi, G., and Cirelli, C. (2011). Sleep and synaptic homeostasis: structural evidence in Drosophila. Science 332, 1576-1581.

27.

Gilestro, G.F., Tononi, G., and Cirelli, C. (2009). Widespread changes in synaptic markers as a function of sleep and wakefulness in Drosophila. Science 324, 109-112.

28.

Vyazovskiy, V.V., Cirelli, C., Pfister-Genskow, M., Faraguna, U., and Tononi, G. (2008). Molecular and electrophysiological evidence for net synaptic potentiation in wake and depression in sleep. Nature neuroscience 11, 200-208.

29.

Liu, Z.W., Faraguna, U., Cirelli, C., Tononi, G., and Gao, X.B. (2010). Direct evidence for wake-related increases and sleep-related decreases in synaptic strength in rodent cortex. The Journal of neuroscience : the official journal of the Society for Neuroscience 30, 8671-8675.

30.

Afonso, D.J., Liu, D., Machado, D.R., Pan, H., Jepson, J.E., Rogulja, D., and Koh, K. (2015). TARANIS Functions with Cyclin A and Cdk1 in a Novel Arousal Center to Control Sleep in Drosophila. Current biology : CB 25, 1717-1726.

113

31.

Allada, R., and Chung, B.Y. (2010). Circadian organization of behavior and physiology in Drosophila. Annu Rev Physiol 72, 605-624.

32.

Abruzzi, K.C., Rodriguez, J., Menet, J.S., Desrochers, J., Zadina, A., Luo, W., Tkachev, S., and Rosbash, M. (2011). Drosophila CLOCK target gene characterization: implications for circadian tissue-specific gene expression. Genes & development 25, 2374-2386.

33.

Fernandez, M.P., Chu, J., Villella, A., Atkinson, N., Kay, S.A., and Ceriani, M.F. (2007). Impaired clock output by altered connectivity in the circadian network. Proceedings of the National Academy of Sciences of the United States of America 104, 5650-5655.

34.

Renn, S.C., Park, J.H., Rosbash, M., Hall, J.C., and Taghert, P.H. (1999). A pdf neuropeptide gene mutation and ablation of PDF neurons each cause severe abnormalities of behavioral circadian rhythms in Drosophila. Cell 99, 791-802.

35.

Hyun, S., Lee, Y., Hong, S.T., Bang, S., Paik, D., Kang, J., Shin, J., Lee, J., Jeon, K., Hwang, S., et al. (2005). Drosophila GPCR Han is a receptor for the circadian clock neuropeptide PDF. Neuron 48, 267-278.

36.

Lear, B.C., Merrill, C.E., Lin, J.-M., Schroeder, A., Zhang, L., and Allada, R. (2005). A G Protein-Coupled Receptor, groom-of-PDF, Is Required for PDF Neuron Action in Circadian Behavior. Neuron 48, 221-227.

37.

Mertens, I., Vandingenen, A., Johnson, E.C., Shafer, O.T., Li, W., Trigg, J.S., De Loof, A., Schoofs, L., and Taghert, P.H. (2005). PDF Receptor Signaling in Drosophila Contributes to Both Circadian and Geotactic Behaviors. Neuron 48, 213-219.

114

38.

Williams, J.A., Su, H.S., Bernards, A., Field, J., and Sehgal, A. (2001). A circadian output in Drosophila mediated by neurofibromatosis-1 and Ras/MAPK. Science 293, 2251-2256.

39.

Cowmeadow, R.B., Krishnan, H.R., and Atkinson, N.S. (2005). The slowpoke gene is necessary for rapid ethanol tolerance in Drosophila. Alcoholism, clinical and experimental research 29, 1777-1786.

40.

Lear, B.C., Lin, J.M., Keath, J.R., McGill, J.J., Raman, I.M., and Allada, R. (2005). The ion channel narrow abdomen is critical for neural output of the Drosophila circadian pacemaker. Neuron 48, 965-976.

41.

Suh, J., and Jackson, F.R. (2007). Drosophila ebony activity is required in glia for the circadian regulation of locomotor activity. Neuron 55, 435447.

42.

Luo, W., and Sehgal, A. (2012). Regulation of Circadian Behavioral Output via a MicroRNA-JAK/STAT Circuit. Cell 148, 765-779.

43.

Jepson, J.E., Shahidullah, M., Lamaze, A., Peterson, D., Pan, H., and Koh, K. (2012). dyschronic, a Drosophila homolog of a deaf-blindness gene, regulates circadian output and Slowpoke channels. PLoS Genet 8, e1002671.

44.

Cavanaugh, D.J., Geratowski, J.D., Wooltorton, J.R., Spaethling, J.M., Hector, C.E., Zheng, X., Johnson, E.C., Eberwine, J.H., and Sehgal, A. (2014). Identification of a circadian output circuit for rest:activity rhythms in Drosophila. Cell 157, 689-701.

45.

Yoshii, T., Wulbeck, C., Sehadova, H., Veleri, S., Bichler, D., Stanewsky, R., and Helfrich-Forster, C. (2009). The neuropeptide pigment-dispersing factor adjusts period and phase of Drosophila's clock. The Journal of

115

neuroscience : the official journal of the Society for Neuroscience 29, 2597-2610. 46.

Yao, Z., and Shafer, O.T. (2014). The Drosophila circadian clock is a variably coupled network of multiple peptidergic units. Science 343, 1516-1520.

47.

Shafer, O.T., Kim, D.J., Dunbar-Yaffe, R., Nikolaev, V.O., Lohse, M.J., and Taghert, P.H. (2008). Widespread receptivity to neuropeptide PDF throughout the neuronal circadian clock network of Drosophila revealed by real-time cyclic AMP imaging. Neuron 58, 223-237.

48.

Fernandez, M.P., Berni, J., and Ceriani, M.F. (2008). Circadian remodeling of neuronal circuits involved in rhythmic behavior. PLoS Biol 6, e69.

49.

Sivachenko, A., Li, Y., Abruzzi, K.C., and Rosbash, M. (2013). The transcription factor Mef2 links the Drosophila core clock to Fas2, neuronal morphology, and circadian behavior. Neuron 79, 281-292.

50.

Petsakou, A., Sapsis, T.P., and Blau, J. (2015). Circadian Rhythms in Rho1 Activity Regulate Neuronal Plasticity and Network Hierarchy. Cell 162, 823-835.

51.

Kula, E., Levitan, E.S., Pyza, E., and Rosbash, M. (2006). PDF cycling in the dorsal protocerebrum of the Drosophila brain is not necessary for circadian clock function. Journal of biological rhythms 21, 104-117.

52.

Kim, E.Y., Bae, K., Ng, F.S., Glossop, N.R.J., Hardin, P.E., and Edery, I. (2002). Drosophila CLOCK Protein Is under Posttranscriptional Control and Influences Light-Induced Activity. Neuron 34, 69-81.

116

53.

Lu, B., Liu, W., Guo, F., and Guo, A. (2008). Circadian modulation of light-induced locomotion responses in Drosophila melanogaster. Genes, Brain and Behavior 7, 730-739.

54.

Kumar, S., Chen, D., and Sehgal, A. (2012). Dopamine acts through Cryptochrome to promote acute arousal in Drosophila. Genes & development 26, 1224-1234.

55.

Zang, Z.J., Gunaratnam, L., Cheong, J.K., Lai, L.Y., Hsiao, L.-L., O'Leary, E., Sun, X., Salto-Tellez, M., Bonventre, J.V., and Hsu, S.I.H. (2009). Identification of PP2A as a novel interactor and regulator of TRIP-Br1. Cellular signalling 21, 34-42.

56.

Sathyanarayanan, S., Zheng, X., Xiao, R., and Sehgal, A. (2004). Posttranslational regulation of Drosophila PERIOD protein by protein phosphatase 2A. Cell 116, 603-615.

57.

Viquez, N.M., Fuger, P., Valakh, V., Daniels, R.W., Rasse, T.M., and DiAntonio, A. (2009). PP2A and GSK-3beta act antagonistically to regulate active zone development. The Journal of neuroscience : the official journal of the Society for Neuroscience 29, 11484-11494.

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Dinis José Silva Afonso - repositorium – Uminho - Universidade do

Dinis José Silva Afonso Regulation of sleep and circadian rhythms by TARANIS Universidade do Minho Escola de Ciências da Saúde Dinis José Silva Afon...

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