The circadian system in insects: Cellular, molecular, and functional organization

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The circadian system in insects: Cellular, molecular, and functional organization Kenji Tomiokaa, Akira Matsumotob a

Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Department of Biology, Juntendo University School of Medicine, Inzai, Japan

b

Contents 1. Circadian rhythms in various physiological functions 2. Clock oscillatory machinery: Insect clock diversity 2.1 Drosophila circadian clock machinery 2.2 Clock machinery in insects other than Drosophila 2.3 Regulation of the molecular clockwork at multiple levels of gene expression 2.4 After effects 3. Circadian organization 3.1 Localization of the circadian clock: Central clocks 3.2 Multicellular organization of the clock 3.3 Peripheral clocks 4. Input pathway 4.1 Photoreceptors 4.2 Entrainment by temperature cycles 4.3 Social influence 5. Output of the clock 5.1 Regulation of behavioural rhythms 5.2 Regulation of sensory systems 5.3 Regulation of learning and memory 5.4 Regulation of photoperiodic time measurement 6. Conclusion and future perspectives: Evolution and diversification of circadian clocks in insects Acknowledgements References

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Abstract The circadian clock organizes the physiology and behaviour of insects to adapt to a daily and seasonally changing environment. The clock oscillates with a period of approximately 24 h in a self-sustained manner, showing an exact 24 h period through synchronization to the daily environmental cycle, and regulates various physiological functions Advances in Insect Physiology ISSN 0065-2806 https://doi.org/10.1016/bs.aiip.2019.01.001

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2019 Elsevier Ltd All rights reserved.

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through neural or humoral pathways. These properties of the clock have been extensively studied at molecular and cellular levels in Drosophila melanogaster since the mid-1980s. During the last 2 decades, progress in molecular biology techniques has promoted studies on the clock system in other insects, including higher and lower phylogenetic groups, such as butterflies, honeybees, crickets, and firebrats, enabling us to compare the system, at least in part, among different insect groups.

Abbreviations AMe BRM CC CK2 Clk cry cwo cyc dbt DD DH31 DN E73 ERG GABA HR3 ITP LD LN LPN luc miRNA OpLW PDF Pdp1 per PG PLC PTTH RDL RNAi tim TTFL loop vri wake β-gal

accessory medulla brahma complex central complex casein kinase 2 Clock cryptochrome clockwork orange cycle double time constant darkness diuretic hormone 31 dorsal neuron ecdysone-induced gene 73 electroretinogram γ-aminobutyric acid hormone receptor 3 ion transport peptide light and dark cycle lateral neuron lateral posteriorly located neuron luciferase microRNA opsin-long wavelength pigment-dispersing factor Par domain protein 1 period prothoracic gland phospholipase C prothoracicotropic hormone resistant to dieldrin RNA interference timeless transcriptional–translational feedback upd unpaired vrille wide awake β-galactosidase

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1. Circadian rhythms in various physiological functions Insects live in harmony with daily cyclic environmental changes, showing diurnal, nocturnal, or crepuscular activity rhythms. For example, swallowtail butterflies visit flowers during the daytime, while cockroaches walk around the kitchen at night, and some female mosquitoes seek host animals in the morning and evening twilight hours. These rhythmic lives are controlled by an endogenous clock mechanism (Saunders et al., 2002). The clock runs with a period, called free-running period, close to, but not exactly, 24 h under constant conditions, hence it is called the circadian clock. It efficiently synchronizes itself to the daily environmental cycles, to show an exact 24-h pattern, mainly by using light or temperature cycles as synchronizing agents. The free-running period is precise and highly resistant to temperature levels and the temperature coefficient Q10 is usually very close to 1.0, thus suggesting that the biochemical mechanism underlying the clock is temperature compensated. However, the mechanism of this temperature compensation is yet to be elucidated. The most important biological significance for the rhythm is the ability to anticipate environmental changes and prepare internal physiological adaptations to face them. The circadian clock plays an important role in various physiological processes; it regulates daily activity rhythms, the timing of mating behaviour, time-of-day-related memory, sun-compass-dependent orientation or navigation, and photoperiodic time measurement for seasonal adaptation (Saunders et al., 2002). Insects are the most prosperous and highly diverged group of animals; they live in various environments, ranging from tropical regions to polar zones and from sea level to high mountains (Engel, 2015; Whitfield and Purcell, 2012). The clock may have played a role in the evolution or diversification of insects. To understand an insect’s evolution/ diversification, elucidation of the circadian clock mechanism, especially its similarity and diversity across various groups of insects, is essential. The underlying mechanism has been studied for several decades and by the end of the 20th century, the basic properties including the neural basis, involved tissues, and the photoreceptors necessary for synchronization to light cycles have been mostly clarified (Saunders et al., 2002). During the last 3 decades, the molecular oscillatory mechanism has been profoundly studied in Drosophila melanogaster, and the basic framework of the clock and its synchronization to the light cycle have been clarified

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(Dubowy and Sehgal, 2017). In this century, molecular and cellular studies have been extended to other insects and deepened our understanding on the commonality and diversity of the clock mechanism as well as the input and output mechanisms (Tomioka and Matsumoto, 2010, 2015). In this review, we will focus on the molecular and cellular bases behind the generation of circadian rhythm, phase regulation of the clock, and the resulting regulatory mechanisms through which the circadian clock controls various physiological functions.

2. Clock oscillatory machinery: Insect clock diversity Studies on the oscillatory mechanism of the circadian clock were initiated in Drosophila during the 1980s and have now been extended to several other insects, although certain aspects still need to be uncovered. In all cases, rhythmic expression of the so-called clock genes is believed to be the core of the clock.

2.1 Drosophila circadian clock machinery The most extensively studied is the Drosophila clock machinery. The basic mechanisms of the clock are transcriptional–translational negative feedback loops, consisting of the so-called clock genes that produce rhythmic gene expression with a period of approximately 24 h (Fig. 1A). One loop consists of the genes Clock (Clk), cycle (cyc), period (per), and timeless (tim) (Tataroglu and Emery, 2015). In this loop, the protein products of Clk and cyc (i.e. CLK and CYC) form a CLK/CYC heterodimer, which activates the transcription of per and tim during late day to early night (Allada et al., 1998; Darlington et al., 1998; Rutila et al., 1998). As transcripts are translated, the proteins PER and TIM accumulate during the night and at late night they form a PER/TIM heterodimer, enter the nucleus, and repress transcriptional activity of the CLK/CYC complex (Lee et al., 1998). The repression reduces the mRNA levels of per and tim, which results in a reduction of their protein products, and releases the CLK/CYC complex from the repression. Then the transcription of per and tim is reactivated and the next round begins. The timing of nuclear translocation and transcriptional repression is controlled through phosphorylation by specific kinases. Double-time (dbt), a casein kinase orthologue, mediates the phosphorylation of PER, which is required for its nuclear entry. It has been shown that dbt mutations affect the timing of nuclear entry (Bao et al., 2001; Price et al., 1998). Casein Kinase 2 (CK2) is also known to phosphorylate PER and regulate the timing

ARTICLE IN PRESS Fig. 1 Molecular oscillatory mechanism of the circadian clock in different insect species: the fruit fly Drosophila melanogaster (A), the monarch butterfly Danaus plexippus (B), and the cricket Gryllus bimaculatus (C). In all cases, the core loop commonly consists of transcriptional activators, CLOCK (CLK) and CYCLE (CYC), and inhibitors PERIOD (PER), TIMELESS (TIM) and/or CRYPTOCHROMEs (CRYs). In addition, other loops regulate rhythmic expression of CLK, CYC and CWO in a species-dependent manner. In B and C, the same loops as Drosophila are shown in grey colour. See text for details.

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of nuclear entry of the PER/TIM complex (Lin et al., 2002). TIM phosphorylation is controlled by shaggy, a glycogen synthase kinase-3 orthologue that consequently regulates the nuclear entry of the PER/TIM complex (Martinek et al., 2001). In addition to the cyclic expression of per and tim, Clk is rhythmically expressed to peak during the day (Fig. 1A). The rhythmic expression of Clk is regulated by a negative element vrille (vri) and a positive element Par domain protein 1 (Pdp1) (Cyran et al., 2003; Glossop et al., 2003; Zheng et al., 2009). Both vri and Pdp1 are transactivated by CLK/CYC complex through E-box during the night. vri instead is transactivated earlier than Pdp1 and the product protein VRI represses the transcription of Clk through V/P-box. Later, when PDP1 accumulates, Clk transcription is triggered by its competitive binding to the V/P-box. Through this mechanism, Clk transcripts and proteins increase during the day. However, more recent studies have shown that PDP1 also acts downstream of the clock to regulate behavioural rhythms (Benito et al., 2007; Lim et al., 2007b). The third loop was found by genome-wide screening of rhythmically expressed genes (Kadener et al., 2007; Lim et al., 2007a; Matsumoto et al., 2007) (Fig. 1A). The constituent gene of this loop is clockwork orange (cwo), a member of the orange superfamily. cwo is also rhythmically expressed by CLK/CYC through E-box, and its product CWO binds to the E-box competitively with CLK/CYC to repress E-box mediated transcription during the day (Zhou et al., 2016). The repression drives highamplitude transcriptional oscillations of the E-box-dependent clock and clock relevant genes.

2.2 Clock machinery in insects other than Drosophila The understanding of how clock machinery works in various insects is slowly improving. The basic machinery is almost the same as Drosophila, including the per/tim negative feedback loop (Fig. 1B and C). However, some variations have been reported. In moths Antheraea pernyi and Bombyx mori, PER is rhythmically expressed in the cerebral neurons, but stays in the cytoplasm and never enters the nucleus (Sauman and Reppert, 1996a; Sehadova´ et al., 2004), while in the beetle Pachymorpha sexguttata and the crickets Teleogryllus commodus and Teleogryllus oceanicus, no rhythmic expression was detected by PER immunohistochemistry (Frisch et al., 1996; Lupien et al., 2003). In the silkmoth A. pernyi, antisense per RNA is suggested to play a role in rhythmic PER expression (Sauman and Reppert, 1996a). In hymenopteran species,

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tim is absent and mammalian-type cryptochrome (cry2) has been speculated to form a feedback loop with per (Rubin et al., 2006; Zhan et al., 2011). The involvement of cry2 in the feedback loop has also been suggested in the monarch butterfly Danaus plexippus by in vitro experiments; CRY2 forms a complex with PER and TIM, enters the nucleus and suppresses the CLK/CYC transcriptional activity (Zhu et al., 2008) (Fig. 1B). cyc is rhythmically expressed to peak during the day in many insects including crickets, bean bugs and firebrats (Ikeno et al., 2008; Kamae et al., 2010; Uryu et al., 2013) (Fig. 1C). In the firebrat, the rhythmic expression of cyc is controlled by ecdysone-induced gene 75 (E75) and hormone receptor gene 3 (HR3) (Kamae et al., 2014). Both genes are thought to be transactivated by CLK/CYC. E75 represses the transcription of cyc and HR3 later re-activates it. In the cricket Gryllus bimaculatus, when cyc is downregulated by RNAi, Clk shows a rhythmic expression (Uryu et al., 2013). This fact suggests that Clk is also under circadian control, but the rhythm is normally inhibited by a cyc-dependent mechanism (Fig. 1C). In addition, cryptochrome (cry) genes are also known to form an oscillatory loop that functions regardless of the per/tim loop in the cricket G. bimaculatus. Like many other insects (Yuan et al., 2007), the cricket has two cry genes, i.e., a Drosophila-type cry (cry1) and a mammalian-type cry (cry2) (Tokuoka et al., 2017). cry2 has several splicing variants and some of the protein products are thought to form a complex with other CRY2 or CRY1 proteins and repress their own transcription by inhibition of CLK/CYC transcriptional activity (Tokuoka et al., 2017) (Fig. 1C).

2.3 Regulation of the molecular clockwork at multiple levels of gene expression In addition to the above described molecular feedback loops, including the posttranslational regulations of clock proteins by phosphorylation and subsequent degradation, recent studies show, mostly in Drosophila, involvement of other factors which influence the molecular clockwork. These factors modulate the abundance of core clock proteins and clock-related proteins through three steps of gene expression: control of initial transcription, post-transcriptional regulation, and translational regulation. In eukaryotes, chromatin remodelling factors are essential for early transcriptional control by changing the degree of chromatin packing, and hence, essential for the access to target sequence elements in the genome by transcription factors. The Brahma complex (BRM) functions catalytically to increase nucleosome density at the promoters of per and tim, creating an

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overall restrictive chromatin landscape that limits transcriptional output during the active phase of cyclic gene expression (Kwok et al., 2015). In addition, the non-catalytic function of BRM regulates the abundance and binding properties of CLK to target promoters and maintains transient RNA polymerase II “stalling” at the per promoter, likely by recruiting repressive and pausing factors. Following transcription, a series of post-transcriptional modifications generally occur, and alternative splicing is one such modification that results in the production of protein isoforms when it occurs in coding regions. When it occurs in non-coding regions, however, the resultant mature transcripts often differ from each other with respect to stability and translational efficiency. In Drosophila clockwork, the latter mechanism was observed after temperature-sensitive splicing in the 30 UTR of per gene (Majercak et al., 2004). At low temperatures, splicing-out of the specific intron dmpi8 in the region leads to the advanced phase of per mRNA and subsequent PER protein accumulation (Table 1). This stimulates the evening peak at the behavioural level. At high temperatures, this splicing-out does not occur and per mRNA accumulates at a slower rate, causing the evening peak of locomotor rhythm to occur at a later phase. The temperature-induced alternative splicing is hypothesized to facilitate adaptation of locomotor rhythm to seasonal changes in day length. The splicing efficiency of the dmpi8 region was recently found to be enhanced by B52/SRp55, a member of the serine/ arginine-rich splicing factors (Zhang et al., 2018) (Table 1). There are ample studies on microRNAs (miRNAs) which modulate abundance and translational efficiency of their target mRNAs at a posttranscriptional level. miRNAs are small non-coding RNAs with a length of about 22 nucleotides (nt). The small RNAs are generated by consecutive cleavage of their precursor transcripts by DROSHA and DICER ribonuclease (Yang and Lai, 2011). In the nucleus, DROSHA cleaves the primary miRNA (pri-miRNA) into a precursor miRNA (pre-miRNA) which is about 70 nt in length. Then the pre-miRNA is exported into the cytoplasm, where DICER cleaves it into miRNA. Subsequently, miRNA loaded into the RNA-induced silencing complex (RISC) recognizes its target mRNA and promotes the target’s degradation. So far, several Drosophila genomic regions have been identified to code miRNAs, which affect various circadian properties like period, phase, and amplitude of clock output (Table 1). In some cases, expression of the miRNA itself is under the control of the circadian clock. For example, miR-263a/b were identified as rhythmically expressed miRNAs in an earlier study (Yang et al., 2008), although little is

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Table 1 Characterization of posttranscriptional regulators which modulates various circadian phenotypes. Expression Factor pattern Target Role of the factor

dmpi8 intron in Phase determination 30 UTR of per

B52/SRp55 miR-263a/b

Rhythmic

miR-276a

Rhythmic

bantam

Not rhythmic 30 UTR of Clk

let-7

Rhythmic

30 UTR of tim

cwo

Inhibition of tim expression Period determination Period determination Phase regulation Morning anticipation

miR-959–964 Rhythmic

Output pathway Innate immunity Metabolism Feeding behaviour

miR-124

Rhythmic

Regulation of BMP signalling pathway Phase regulation of evening peak

miR-279

Not rhythmic upd

miR-996 miR-92 LARK

Regulation of JAK/STAT signalling pathway Rhythm generation

SIRTUIN 2

Modulation of neuronal excitability of PDF cells

Not rhythmic dbt-RC and Specific promotion of dbt-RC dbt-RE isoforms translation Translation of dbt-RE induced by light Rhythm generation

TYF

Not rhythmic per

Interaction with ATX2 Rhythm generation

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known about how they are regulated. In the case of miR-276a, detailed studies have revealed that it is rhythmically induced by the transcription factor Chorion factor-2 under conditions of light, and it targets the 30 UTR of tim (Chen and Rosbash, 2016). bantam is the first miRNA whose role was clarified in the circadian system (Kadener et al., 2009) (Table 1). Over-expression of bantam in pacemaker neurons lengthens the circadian period through regulation of an interaction between RISC and the target sites of bantam, which is found in 30 UTR of Clk mRNA (Kadener et al., 2009). Thus, bantam is postulated to affect circadian period via controlling the abundance of CLK. let-7 is the other miRNA with the capacity to lengthen circadian period when it is overexpressed in neurons that express the neuropeptide pigment-dispersing factor (PDF) (Chen et al., 2014) (Table 1). let-7 knockout, in turn, causes weakening of locomotor rhythmicity, which eliminates the morning anticipation. Interestingly, one of the let-7 targets is cwo, and let-7 ablation induces faster accumulation of PER and mis-projection of PDF neurons (Chen et al., 2014). Output pathways of the circadian clock are also targeted by miRNAs (Table 1). miR-959–964 are rhythmically expressed, peaking at early night, as cluster miRNAs derived from the same primary-miRNAs (Vodala et al., 2012). These six miRNAs are mainly expressed in the fat bodies of the adult head. Results of cluster knockout and overexpression experiments suggest that these miRNAs regulate specific circadian output pathways, innate immunity, metabolism, and feeding behaviour. The other example can be seen in miR-124. Its knockout leads to an advance of evening peak of the locomotor rhythm in light/dark cycle (LD) with no effect on the circadian period under constant darkness (DD) (Garaulet et al., 2016; Zhang et al., 2016). Because this advanced evening peak was rescued by inhibition of the bone morphogenetic protein (BMP) signalling pathway (Garaulet et al., 2016), miR-124 is assumed to regulate diverse aspects of rhythmic behaviour through the BMP pathway. JAK/STAT pathway is identified as the other clock output signalling pathway regulated by miRNAs, miR-279 and miR-996 (Sun et al., 2015) (Table 1). Both over-expression and knockout of miR-279 abolished locomotor rhythms, and this miRNA targets unpaired (upd) in the JAK/STAT signalling pathway (Luo and Sehgal, 2012). Thus, cells expressing both miR-279 and upd are thought to form the physical connection between pacemaker neurons and output cells, generating rest-activity rhythms. Neuronal excitability of PDF positive pacemaker neurons is also known for the

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target of miRNAs. This is mediated by miR-92a, which is the only nocturnal miRNA found in PDF positive cells (Table 1). Chen and Rosbash (2016) reported that miR-92a modulates PDF neuronal excitability through suppressing SIRTUIN 2, which belongs to the Sirtuin family having an NAD-dependent deacetylase. The clock information is then transmitted to other cells with PDF (Hyun et al., 2005), and the signalling pathway downstream of the PDF receptor is regulated by the miRNA GW182, through silencing the expression of the cAMP phosphodiesterase DUNCE in a light-dependent manner (Zhang and Emery, 2013). Recent studies have also elucidated the importance of translational regulation in the circadian timekeeping mechanism. Many clock-regulated transcripts are synchronously translated at either midday or midnight in Drosophila pacemaker neurons (Huang et al., 2013). Translation of core clock genes is also specifically regulated at a translational level. Double-time (dbt) is one of such translationally regulated genes. Translation of its transcripts are directly regulated by LARK, a rhythmic RNA-binding factor, which promotes translation of specific alternative dbt transcripts, called dbt-RC and dbt-RE (Huang et al., 2014) (Table 1). Interestingly, the translation of the latter isoform is induced by light in a lark-dependent manner. Another example of translational regulation is per, since its translation is activated by the product protein of the twenty-four gene (TYF) (Table 1). TYF interacts with ATAXIN-2, a RNA-binding protein, to coordinate an active translation complex that is important for translation of per transcripts (Lee et al., 2017; Lim et al., 2011; Lim and Allada, 2013; Zhang et al., 2013).

2.4 After effects It has been known that some properties of the circadian clock in constant darkness are modulated or determined by light conditions preceding constant darkness. For example, the free-running period of the cockroach locomotor rhythms in DD is shorter when entrained to LD11:11 h than when entrained to LD13:13 h, and the period persists for a long time (Page, 1983a). Similar modulation of the free-running period is reported for Drosophila where different photoperiods in a 24 h light cycle, applied during larval and pupal development, significantly affected the free-running period in the following DD (Tomioka et al., 1997). In the cricket, both free-running periods and duration of the active phase in DD were determined by photoperiods of the preceding 24 h light cycle (Koga et al., 2005; Tomioka and Chiba, 1989a, 1989b). The effects were also detected in neuronal activity rhythms in the

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cricket optic lobe (Koga et al., 2005; Tomioka and Chiba, 1989a) and could be transferred to another individual by optic lobe transplantation in the cockroach (Page, 1982). Therefore, these results seem to confirm that regulation occurs in the clock tissue. The molecular mechanism underlying the aftereffects remains largely unknown, but there is a possibility that epigenetic regulation is involved. Chromatin modification may be involved in the after-effect; in crickets, methylation on H3K27 by Enhancer of zeste is required for photoperiodic modulation of the active phase duration and free-running period, which is associated with changes in expression profiles of clock component genes (Hamada et al., 2016).

3. Circadian organization 3.1 Localization of the circadian clock: Central clocks The clocks have been localized in several insect species in respect to those controlling overt activity rhythms. In hemimetabolous insects, including crickets and cockroaches, the optic lobe is the locus of the clock that regulates the activity rhythm (Page, 1988; Tomioka, 2014). The optic lobe was first postulated as the candidate tissue by a surgical lesioning experiment in the cockroach Leucophaea maderae (Nishiitsutsuji-Uwo and Pittendrigh, 1968a). Similar results were reported in crickets (Loher, 1972; Tomioka and Chiba, 1984). The clock site was further defined by partial removal of the optic lobe in the cockroach and the ventral proximal region was reported to be the likely locus of the clock (Page, 1978; Roberts, 1974; Sokolove, 1975). However, the lesioning experiments did not provide crucial evidence for the clock site, because the lesion may just destroy the output pathway of the clock, not the clock itself. To define the clock locus unequivocally, transplantation and electrophysiological experiments were subsequently performed. Optic lobe transplantation in the lobotomized cockroach restored the rhythm with characteristics of the donor’s rhythm, providing strong evidence for the optic lobe as the clock locus (Page, 1982, 1983b). A similar result was also reported in the cricket (Tomioka, 2014). Unequivocal evidence that the clock is localized in the optic lobe was given by recording the neuronal activity from the tissue isolated from the nervous system in situ or in vitro. The isolated optic lobe maintained the neuronal activity rhythms under constant conditions both in crickets and cockroaches (Colwell and Page, 1990; Tomioka and Chiba, 1986, 1992). In holometabolous insects, including moths, mosquitoes, and flies, the optic lobe removal did not prevent the activity rhythms, suggesting that

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the clock is located outside the optic lobe (Helfrich et al., 1985; Kasai and Chiba, 1987; Truman, 1974). In Drosophila, genetic ablation of the optic lobe suggests that this is dispensable for the circadian locomotor rhythms (Helfrich and Englemann, 1983). Transplantation experiments further showed that the clock is located in the central brain in moths and Drosophila (Handler and Konopka, 1979; Truman, 1971). The clock locus in these insects has been later confirmed by detection of clock gene products. For example, the protein product of per can be detected by immunohistochemistry using anti-PER antibodies. In D. melanogaster, PER is rhythmically expressed in about 150 cerebral neurons located in specific regions (Hermann-Luibl and Helfrich-F€ orster, 2015; Siwicki et al., 1988). In the moth A. pernyi, the PER immunoreactivity is localized in cells of the pars intercerebralis (Sauman and Reppert, 1996a). Taken together, the central clock regulating overt activity rhythms is located in the protocerebrum, although the exact location differs among phylogenetic groups. It is also a general feature that the clock consists of bilaterally paired structures.

3.2 Multicellular organization of the clock Immunohistochemistry using antibodies against the clock gene products or in situ hybridization using DNA or RNA probes of the clock genes revealed that multiple cells express the clock genes in the identified clock tissues (Fig. 2). For example, many cells in the optic lobe express per and cry2 in G. bimaculatus (Kutaragi et al., 2018) (Fig. 2D). Immunohistochemistry revealed that several cells in the optic lobe and central brain express PER in the cockroach Blattella germanica (Wen and Lee, 2008) (Fig. 2B) and in the crickets, T. commodus and T. oceanicus (Lupien et al., 2003). The cells expressing TIM are located in several sites of the protocerebrum, including the lobula, pars intercerebralis, and pars lateralis in the monarch butterfly (Zhu et al., 2008) (Fig. 2C). These results suggest the multicellular organization of the circadian clock. The cellular organization of the cerebral clock has been extensively analysed in D. melanogaster. There are seven groups of clock neurons bilaterally paired, forming a dynamic network in the Drosophila brain, i.e., large and small ventral lateral neurons which are located in the ventral region between the optic and the protocerebral lobes (lLNv, sLNv), dorsal lateral neurons (LNd), three groups of dorsally located neurons (DN1, DN2, DN3), and lateral posteriorly located neurons (LPN) (Hermann-Luibl and

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A

D

B

C

Fig. 2 Schematic diagrams showing clock gene or clock protein expressing cells in the €rster, 2002), the cockroach Blattella germanica fruit fly D. melanogaster (A) (Helfrich-Fo (B) (Wen and Lee, 2008), the monarch butterfly D. plexippus (C) (Zhu et al., 2008), and the cricket G. bimaculatus (D) (Kutaragi et al., 2018). (A) In Drosophila, there are seven main groups of neurons. Four groups (LNd, lLNv, sLNv, and LPN) are located laterally, and the remaining three groups (DN1, DN2, and DN3) are located in the dorsal region. The sLNv and lLNv express a neuropeptide, PDF, except one sLNv (5th sLNv) expressing ITP. (B) In Blattella, three groups of PER immunoreactive neurons (PER-IN) are located in the dorsal protocerebrum (D1, D2, and D3). Three groups of PER-IN located in the optic lobe and one group in the deutocerebrum coexpress PDF (PDFMe, PDFLad, PDFLav, and PDFDe). (C) In Danaus, TIM immunoreactive cells are located in the lobula (LO), pars lateralis (PL), pars intercerebralis (PI), and subesophageal ganglion (SOG). (D) In Gryllus, per and cry2 expressing neurons are located near outer chiasma between lamina and medulla in the optic lobe. A–C, frontal view. D, dorsal view. OL Optic lobe, Pr protocerebrum, De deutocerebrum, La lamina, Me medulla, Lo lobula. For further explanations see text.

Helfrich-F€ orster, 2015; Kaneko, 1998; Yoshii et al., 2005) (Fig. 2A). They are characterized by the size and location of their cell bodies as well as by their different neurotransmitters. The sLNv and lLNvs, except for 5th sLNv, express PDF. Anteriorly located DN1a neurons express a CCHa1 peptide (Fujiwara et al., 2018), while posteriorly located DN1p neurons express a diuretic hormone 31 (DH31) (Kunst et al., 2014). The 5th sLNv and one

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LNd cell are characterized by an ion transport peptide (ITP) (HermannLuibl et al., 2014). In addition, recent cellular transcriptome analyses revealed that the phase of clock gene expression, as well as expression pattern of clock controlled genes, are quite different among LNv, LNd, and DN1 (Abruzzi et al., 2017). Thus, the clock is a network consisting of heterogeneous groups of clock neurons that have different properties, hence they play different roles in the clock cellular network. Similar cellular organization of the cerebral clock was found in different Drosophila species: the clock consists of lateral and dorsal neuron clusters (Hermann et al., 2013). The expression of CRY and PDF was found to be species-specific, while ITP was consistently expressed in the 5th sLNv and one LNd cell. Thus, at least among Drosophila species, the cellular organization of the clock is highly conserved, and ITP probably plays a common role in the clock cellular network across species while PDF might be involved in species-specific roles. The importance of multicellular organization was recently stressed in a study using dispersed cell culture of clock neurons in D. melanogaster (Sabado et al., 2017). Although CLK/CYC-mediated transcription was constantly active in dissociated clock neurons, only 10% or less of the cells show circadian expression of PER. In most cells, PER and TIM are abundantly expressed in the cytoplasm but no nuclear accumulation of PER is observed. Given the fact that electrical activity of clock neurons are necessary for molecular oscillation (Nitabach et al., 2002), Drosophila clocks seem to be weak oscillators and need to interact through their network to generate robust 24 h rhythms.

3.3 Peripheral clocks In many insects, circadian clocks are now recognized to reside in various tissues other than the central clock tissue. The peripheral clock was first shown in isolated and incubated salivary glands in D. melanogaster (Weitzel and Rensing, 1981). The isolated glands showed rhythmicity in membrane potential detected by the fluorescent dye, 3,30 -dihexyl-oxacarbocyanine iodide. Circadian rhythms were later detected in various peripheral tissues by a clock gene reporter assay. In the first experiments, the Malpighian tubules were shown to harbour a brain-independent circadian clock, using β-gal as a reporter for per expression (Giebultowicz et al., 2000; Hege et al., 1997). Subsequently a per-luciferase reporter was used, and a wide variety of tissues were detected to show circadian rhythms-independent of the cerebral

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central clock (Plautz et al., 1997). Similar evidence has been reported for other insects, including moths, crickets, and cockroaches (Merlin et al., 2007, 2009; Uryu and Tomioka, 2010; Wiedenmann et al., 1986). These facts suggest that the insect’s whole body is the multicellular clock system in which constituent clocks are somehow temporally organized. The temporal organization of the system will be discussed later.

4. Input pathway 4.1 Photoreceptors Synchronization, or entrainment by daily environmental cycle, is an important property of the circadian clock to set behavioural and physiological events to occur at an appropriate time of day. Light is the most reliable “time cue” or “zeitgeber” for this entrainment. It resets the phase of the circadian clock in a time-of-day-dependent manner: a light pulse causes an advance or a delay phase shift when given at late subjective night or early subjective night, respectively, while it induces little shifts during the subjective day ( Johnson, 1990). The clock with a period longer than 24 h synchronizes to the 24 h light cycle by an advance shift caused by an exposure of late subjective night to morning light, while the clock with a period shorter than 24 h achieves synchronization by an exposure of early subjective night to evening light. The photoreceptor necessary for the photic entrainment has been identified in several insects. The most well-studied one is the cryptochrome (CRY: dCRY or CRY1) of Drosophila. CRY1 is a member of the photolyase superfamily and comes into play in clock resetting as a blue light receptor in several cerebral clock neurons. It is activated by blue light and leads to TIM degradation by cooperation with JETLAG, an F-box protein with leucine-rich repeats (Koh et al., 2006; Peschel et al., 2009) (Fig. 3A). TIM is rhythmically expressed in the clock neurons, increasing during the night, and its degradation causes either phase delay at accumulating phase (early subjective night) or phase advance at decreasing phase (late subjective night). A similar mechanism has been suggested in the monarch butterfly, D. plexippus (Zhu et al., 2008). The compound eye is widely used for synchronization among insects, including Drosophila. Hemimetabolous insects use the compound eye almost exclusively for photic entrainment. Thus, bilateral sectioning of the optic nerve prevents photic entrainment of the rhythm (Nishiitsutsuji-Uwo and Pittendrigh, 1968b; Tomioka and Chiba, 1984). Only in a few cases has the detailed entrainment mechanism been studied. In the cricket,

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Fig. 3 Light entrainment mechanisms of the circadian clock. (A) Cryptochrome (CRY1)dependent resetting of the clock in D. melanogaster and D. plexippus. Light-activated CRY1 leads TIM degradation through a mechanism involving JETLAG (JET). (B) Compound-eye-dependent resetting of the clock in G. bimaculatus. Light information perceived by green-sensitive opsin (OpLW) acts on the clock cell by neurotransmitters which activate the Pdp1 pathway and/or c-fos pathway. Elongation of light phase upregulates Pdp1 (PDP1) which increases CLK then per and tim to prolong the night phase, while c-fosB (C-FOSB) upregulated by light resets the CRY-loop through posttranscriptional regulation of crys (CRY). See text for detailed explanations.

G. bimaculatus, the compound eye is the necessary circadian photoreceptor and green-light sensitive opsin (opsin-long wavelength, OpLW) is the major receptor molecule (Komada et al., 2015). RNAi knockdown of the opLW gene severely affected the photic entrainment. The photic information perceived by OpLW is transmitted to the optic lobe and resets the phase of the

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clock, which is most likely located near the outer chiasma, between lamina and medulla. There are at least two intracellular pathways for resetting (Fig. 3B). One is the Pdp1-dependent pathway-induced by a delayed termination of light phase: prolonged light exposure upregulates level of Pdp1 mRNA, which is followed by Clk, per and tim upregulation (Kutaragi et al., 2016). This prolongs the subjective night phase, and thus delays the clock oscillation. The second pathway includes c-fosB and two cry genes, cry1 and cry2 (Kutaragi et al., 2018). Light information up-regulates c-fosB expression, after which a pathway involving the cry genes is activated. In crickets, RNAi of either cry1 or cry2 does not prevent photic entrainment of the treated crickets (Tokuoka et al., 2017). Therefore they are not circadian photoreceptors, but play a role in resetting the clock through a negative feedback loop that they form, and that can operate independently of the per/tim loop (Tokuoka et al., 2017). Although detailed mechanisms through which light resets this loop need to be elucidated, the resetting of cry loop probably influences the per/tim loop through CLK/CYC as a hinge. The neurotransmitter involved for this resetting mechanism remains unidentified, but GABA and Mas-allatotropin are the candidates, since they are known to cause phasedependent phase shifts of locomotor rhythm which are similar to those caused by light in L. maderae (Petri et al., 2002).

4.2 Entrainment by temperature cycles The temperature cycle is also an important zeitgeber to synchronize the circadian clock. Although there is ample evidence for the entrainment of circadian clocks to temperature cycles, the mechanism is not well understood. In Drosophila, the temperature is perceived through a nocte gene mediated mechanism by a receptor in the chordotonal organ located in the thorax (Glaser and Stanewsky, 2005; Sehadova et al., 2009). The information is conveyed to the cerebral clock neurons through a neural pathway and most likely resets the circadian oscillation in LNd and DNs to synchronize to the temperature cycle (Chen et al., 2018; Miyasako et al., 2007; Picot et al., 2009). Temperature cycles can entrain the oscillation through upregulation or downregulation of clock gene expression in a temperature-dependent manner, with Clk as the first responder (Yoshii et al., 2005, 2007). Mutant flies for the norpA gene, which encodes for the enzyme phospholipase C (PLC), cannot synchronize to temperature cycles (Glaser and Stanewsky, 2005), suggesting that PLC is involved in temperature perception or in signal transduction in the temperature entrainable clock neurons.

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In other insects, behavioural entrainments by temperature cycles are wellknown, but the underlying molecular mechanisms are still poorly understood. In the cricket G. bimaculatus, the temperature entrainment mechanism was examined with respect to the rhythmic expression of clock genes. When temperature cycles were shifted by 6 h under constant darkness, the expression rhythm of per and tim rather quickly resynchronized to the temperature cycle while cyc did slowly with more transient cycles (Kannan et al., 2019). This fact suggests that temperature-sensitivity or temperatureentrainability differs among clock genes. Temperature cycles can also cause rhythms through a mechanism different from that involving the central clock. In Drosophila, per0 arrhythmic mutant flies show a clear locomotor rhythm under temperature cycles, and the phase angle between the temperature cycle and the activity onset systematically changes depending on the period of the given temperature cycle (Yoshii et al., 2002). A similar finding was reported in the cricket T. commodus: optic lobotomized crickets showed a stridulatory activity rhythm under temperature cycles only with a period close to 24 h (Rence and Loher, 1975). These facts indicate that apart from the central clock, a temperature-entrainable circadian clock is also involved; however, its entity as well as oscillatory mechanism are yet to be elucidated.

4.3 Social influence Most insects in nature live with other conspecific or heterospecific individuals. The interaction often affects circadian rhythms of individuals or populations (Bloch et al., 2013). Honey bees are typical eusocial insects that live in a colony, forming a social community where individuals play different roles. Worker bees show a clear circadian rhythm which synchronizes with the individuals of the colony. The synchronization is achieved by the influences from the workers and the queen (Moritz and Sakofski, 1991; Southwick and Moritz, 1987) which stably entrain the clock even in a situation with conflicting light and social environmental cues (Fuchikawa et al., 2016). Although the key factor(s) involved in the synchronization still remains to be elucidated, direct contact between bees and indirect influences through temperature and pheromones are suggested to be involved (Moritz and Kryger, 1994; Southwick and Moritz, 1987). The influence of interaction between individuals on the circadian rhythm is also seen in non-social insects. In D. melanogaster, when a male and a female are housed in a small chamber, their locomotor rhythms and

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daily sleep rhythms change drastically (Fujii et al., 2007; Kawaguchi et al., 2016). These changes are associated with changes in molecular oscillation in the cerebral clock neurons (Fujii and Amrein, 2010; Hanafusa et al., 2013). This social interaction requires an olfactory receptor, Or47b expressed in antennae (Fujii et al., 2007; Lone and Sharma, 2012), which detects three compounds of fly-derived fatty acid methyl esters (Dweck et al., 2015).

5. Output of the clock The circadian clock regulates various overt rhythms, including activity rhythms, circadian changes in memory formation and recall, timecompensated sun-compass navigation, and rhythmic changes in sensitivity of sensory systems. The regulatory mechanisms underlying these overt rhythms have been studied at cellular and molecular levels.

5.1 Regulation of behavioural rhythms 5.1.1 Ecdysis and eclosion rhythms Eclosion, or adult emergence from pupae, is a once-in-a-lifetime event, but certainly controlled by the circadian system. In the silkmoths A. pernyi and Hyalophora cecropia, the brain determines eclosion timing at late or early day, respectively, through a humoral pathway; this has been shown by brain transplantation experiments between the two species (Truman, 1971). Involvement of two hierarchically organized circadian clocks in the underlying circadian system has been shown in the fruit fly, Drosophila pseudoobscura (Pittendrigh et al., 1958; Tackenberg et al., 2017). A master clock (A-oscillator) synchronizing to the light cycle entrains a slave clock (B-oscillator) that regulates eclosion behaviour (Fig. 4A). The B-oscillator is insensitive to light but not to temperature and shifts its phase in response to temperature changes. When a light pulse is given during the subjective night, the A-oscillator is instantaneously reset by the pulse, while the B-oscillator gradually shifts its phase to regain the original phase relationship with the A-oscillator. The process of resynchronization of the B-oscillator is observed as transient cycles. Physiological and molecular studies have deepened the knowledge regarding possible A- and B-oscillators in D. melanogaster: the A-oscillator is probably located in LNvs and the B-oscillator in the prothoracic gland (PG) (Selcho et al., 2017). When LNvs are genetically ablated, both the eclosion rhythm and molecular oscillation of TIM in the gland are diminished (Myers et al., 2003). PER oscillation in the PG clock receives the light information

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B

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Fig. 4 Multioscillator/clock organization regulating overt rhythms. (A) A–B twooscillator model in Drosophila pseudoobscura. Light entrainable A oscillator governs temperature-sensitive B oscillator which drives eclosion behaviour. (B) L–R two clocks regulating locomotor rhythm in crickets and cockroaches. Bilaterally paired left (L) and right (R) clocks mutually synchronize and inhibit output during subjective days to produce stable nocturnal activity. (C) M–E two clocks regulating locomotor rhythms in D. melanogaster. Under LD at constant temperature, both M- and E-clocks synchronize to LD and regulate the morning and the evening peak, respectively. When exposed to LD and temperature cycles simultaneously, the M- and a part of E-clock neurons synchronizing to LD determine active phase, drive morning peak, and govern temperature entrainable E-clock neurons through PDF. Open arrows indicate entraining signal from LD or temperature cycles. Dotted arrows indicate internal entrainment signal between clocks. See text for detailed explanations.

from the central clock, and the control from the LNs is important to maintain the robust oscillations in the PG (Morioka et al., 2012). The timing of ecdysis is also a unique event precisely controlled by hormones. A brain neurohormone, prothoracicotropic hormone (PTTH), is released at a specific time, triggering a series of physiological events that lead to ecdysis (Tomioka and Bollenbacher, 1989; Truman and Riddiford, 1974). PTTH is produced in cells of the pars lateralis in the protocerebrum and secreted from their terminals in the neurohaemal organ, corpora allata in the silkmoth A. pernyi (Sauman and Reppert, 1996b). The hormone stimulates the synthesis of ecdysone in the PG which also harbour the clock for timed ecdysone release (Mizoguchi and Ishizaki, 1982). Thus, this system also represents the hierarchically organized A–B two oscillator systems. The synthesized and released ecdysone regulates gene expression of cells in various tissues to cause metamorphosis through ecdysis and eclosion at an appropriate timing.

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5.1.2 Locomotor rhythm Locomotor rhythms are the most prominent overt rhythm controlled by the clock. The regulatory pathway is principally neuronal in crickets and cockroaches because these insects lose the overt locomotor rhythms when optic tracts are severed or when the optic lobes are bilaterally removed. Lobectomized, hence arrhythmic, cockroaches and crickets restore the rhythm when the optic lobes are transplanted and their neural connection to the protocerebral lobe is regenerated (Page, 1982, 1983b; Tomioka, 2014). This pathway probably uses PDF as a neurotransmitter for regulating, at least in part, the locomotor activity. PDF is produced and stored in the cells located in the proximal medulla area and in those located in the dorsal and ventral regions of the outer chiasma, between lamina and medulla (Fig. 2B). Those cells are called PDFMe, PDFLad, and PDFLav, respectively (Abdelsalam et al., 2008). The processes of PDFLad and PDFLav innervate the lamina area, and those of PDFMe innervate the whole medulla area and their axons project toward the central protocerebrum (Abdelsalam et al., 2008; Okamoto et al., 2001). When PDF content was reduced by pdf RNAi, the locomotor activity was severely suppressed in crickets and the rhythm was diminished in the German cockroach (Hassaneen et al., 2011; Lee et al., 2009). The clock system driving the locomotor rhythm consists of bilaterally paired clocks in crickets and cockroaches, in which each of the two optic lobes harbours a clock (Fig. 4B). The right and left clocks are probably equivalent and coupled to form a pacemaker which produces a free-running period slightly shorter or longer than that of a single clock (Okada et al., 1991; Page et al., 1977). The coupling is achieved by exchanging the light and circadian information through a neuronal pathway in which serotonin may be involved as a neurotransmitter (Tomioka, 1999; Yukizane and Tomioka, 1995). The two clocks also mutually inhibit their output signal during the subjective day to enhance nocturnality (Page, 1983a; Tomioka et al., 1991). The clock system controlling locomotor rhythms has been profoundly studied at cellular and molecular levels in D. melanogaster. As mentioned above (Section 3.2), the central clock consists of seven groups of clock neurons (Fig. 2A). The principal cells for controlling locomotor rhythms are PDF expressing LNvs, because flies lacking PDF or PDF-expressing cells showed abnormality of locomotor rhythms with no morning peak and an earlier evening peak under LD, and short period rhythms that quickly dampen in DD (Lin et al., 2004; Renn et al., 1999). Molecular genetic

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studies revealed that these PDF-expressing LNvs control the morning peak, hence called M-cells, and the LNd, DNs and PDF-lacking sLNv (5th sLNv) control the evening peak, hence called E-cells (Grima et al., 2004; Rieger et al., 2006; Stoleru et al., 2004) (Fig. 4C). PDF expressing LNvs are necessary, and rescue of clock functions in these cells is sufficient for freerunning locomotor rhythm in DD and morning anticipation in LD (Grima et al., 2004; Stoleru et al., 2004). These groups of cells have characteristics for light and temperature “entrainability”. The M-cells and some of the E-cells are expressing CRY and are predominantly light- entrainable, while the other E-cells are temperature-entrainable (Fig. 4C). These properties have been elucidated by entrainment experiments by combined light and temperature cycles in both locomotor rhythms and molecular cycling of clock gene protein products (Miyasako et al., 2007; Yoshii et al., 2009). With this dual clock system, flies can be active at appropriate times of a day in natural field conditions. The M-cells firmly synchronize to the light cycle and normally regulate the phase of E-cells by the PDF signalling pathway, because in pdf 01 flies lacking PDF the phase of evening peak occurs even more advanced than control when the temperature cycle was advanced relative to the LD (Miyasako et al., 2007). In this context, the M–E clock system is in part similar to the A–B two oscillator system proposed by Pittendrigh et al. (1958). The dominance of the M-cells over E-cells is also shown by genetic manipulation of the free-running period (Stoleru et al., 2005). Under constant darkness, the pace of the M-cells can govern the free-running period of the E-cells while the manipulation of the period of E cells does not affect the M-cells. However, the hierarchical master-slave organization is not rigidly fixed, but flexibly changes when confronting various environmental conditions. In short-day conditions, the M-cells indeed dominate and govern the pace of the E-cells. In long-day conditions, however, the E-cells works as a master clock to govern the free-running period of the M-cells (Stoleru et al., 2007). Thus, the circadian clock network property is dynamically changeable to optimize the rhythm in various situations. Beside these clock systems with circadian clock neurons, an ultradian oscillator based mechanism is proposed for the circadian control of locomotor rhythms in the cockroach L. maderae (Schneider and Stengl, 2005). In L. maderae, the accessory medulla (AMe) of the optic lobe is thought to be the locus of the circadian clock (Reischig and Stengl, 2003). The cells form assemblies in the AMe and those in the same assembly fire synchronously

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at regular intervals by a mechanism mediated by GABAergic inhibition. The firing frequency differs among assemblies but is unified by PDF to gate circadian outputs to control the locomotor rhythm. There are several studies showing that the locomotor rhythm is regulated by a humoral pathway. These are shown by brain transplantation experiments: e.g., the fruit fly D. melanogaster and the cricket Acheta domesticus, in which locomotor rhythm was restored after transplantation (Cymborowski, 1981; Handler and Konopka, 1979). In these cases, some neuroendocrine or hormonal factor(s) must be involved in the regulation of locomotor rhythms since the restoration of the rhythm occurred almost immediately after the transplantation. However, the humoral factors have yet to be elucidated. 5.1.3 Sleep-arousal rhythms Sleep-like behaviour has been known for long time in insects (Andersen, 1968; Kaiser and Steiner-Kaiser, 1983). However, the sleep was later defined as a quiescent state, characterized by a decreased responsiveness to sensory stimuli, distinguishable from coma or anaesthesia by its quick reversibility with a sufficiently strong stimulus, and its timing is regulated by the circadian clock (Dubowy and Sehgal, 2017). The circadian regulation of sleep has been extensively studied in Drosophila (Dubowy and Sehgal, 2017). lLNvs play important roles in this mechanism by promoting arousal of flies with their firing activity, which is controlled by light and GABA. Under lightdark cycles, their firing rate fluctuates over the course of the day, increasing around dawn and late in the day in association with arousal (Liu et al., 2014; Shang et al., 2008). The GABAergic sleep-promoting neurons suppress firing in both lLNvs and sLNvs through a GABA-A receptor, Resistant to dieldrin (RDL), to promote sleep (Chung et al., 2009; Parisky et al., 2008). This GABAergic sleep promotion is probably mediated by wide awake (wake) expressed in lLNvs (Liu et al., 2014). The expression of wake is under circadian control and is upregulated to peak at dusk. The increased levels of its product protein WAKE upregulates RDL GABA-A receptor, enhancing sensitivity to GABA. The enhanced sensitivity to GABA then results in suppression of lLNv activity, hence in the promotion of sleep after lights-off. Alternatively, a reduction of GABAa receptors by CLKdependent expression of Fbxl4 increases the lLNv activity and promotes wakefulness (Li et al., 2017). Besides lLNvs, DN1 clock neurons also play an important role in sleep regulation through the diuretic hormone 31 (DH31) as their signal molecule (Kunst et al., 2014). Knock-down of DH31 increases sleep during late night,

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while its overexpression in these cells reduces sleep. Activation of PDF receptor in DH31-expressing DN1s suppresses sleep during this period, but has no apparent effects on the free-running locomotor rhythm, suggesting that this DH31 signalling is downstream of PDF and a part of the sleep specific circadian output pathway to promote wakefulness (Kunst et al., 2014). Interestingly, however, DN1s have a role in the promotion of sleep during midday and night-time (Guo et al., 2016). In this case, they use glutamate as their signal molecule. Optogenetic or thermogenetic activation of glutamatergic DN1 clock neurons increases daytime sleep and suppresses the evening activity peak. In contrast, silencing those neurons reduces sleep during early night. The mechanism by which the switching between the two opposing roles in DN1s is achieved needs to be elucidated. In insects other than Drosophila, sleep has not been extensively studied, and only a few reports exist on this topic. However, many insects should show sleep-like behaviours as noted above. Thus, sleep mechanism and its circadian control should be an important theme to understand daily behavioural adaptation and its diversification among insects. 5.1.4 Navigation The circadian clock plays an important role in time-compensated sun compass (or sky compass) orientation which is known to be pivotal for several insects, including butterflies, honey bees, locusts, and ants (Hoinville and Wehner, 2018; Homberg, 2015; Lindauer, 1960; Reppert et al., 2016). Among them, the most extensively studied is the monarch butterfly D. plexippus, which shows yearly long-distance migration in North America. In fall, butterflies migrate from their breeding area in northern America to overwintering sites in central Mexico, where they congregate on the oyamel fir trees of the small area in the States of Mexico and Michoacan and remain there until spring. In spring, the overwintered butterflies begin to return northward for reproduction and their offspring continue to migrate northward to expand the habitat range (Reppert et al., 2016). During both the southward and the northward migration, the monarch butterflies utilize the time-compensated sun compass supported by the circadian clock (Guerra and Reppert, 2015; Reppert et al., 2016) (Fig. 5). An important information for the orientation is daylight cue, which is perceived by the compound eyes. The main retina senses the azimuthal position of the sun as the dominant source of directional information (Mouritsen and Frost, 2002), while the polarization pattern of UVs is also sensed by UV-photoreceptors in the dorsal rim area of the compound eyes

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Skylight input

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Polarization pattern (Dorsal rim) PL

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Fig. 5 Time-compensated sun compass in the monarch butterfly D. plexippus. The sunlight is perceived by compound eyes, cerebral clock neurons, and antennal clock cells. The polarized light information perceived by the dorsal rim area of the compound eye, and the temporal information from the antennal clock, are integrated into the central complex (CC) to compensate the sun compass, enabling the butterfly to orient accurately to the destination. The brain clock in the pars lateralis (PL) is thought to be less important for this process.

(Labhart et al., 2009; Reppert et al., 2004). Because the sun’s position in the sky continuously changes throughout a day, the flight direction of the butterfly must be adjusted according to the skylight cues, the sun’s position, and the polarization pattern, depending on the time of day. Interestingly, circadian clocks in the antennae are important to yield the light-entrained time

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information (Merlin et al., 2009; Sauman et al., 2005; Shlizerman et al., 2016). The clock neurons are also identified in the pars lateralis by immunohistochemistry, but their effect on the sun compass orientation is thought to be rather minor (Reppert et al., 2016). The time information from antennal clocks is connected to the main sun compass cells existing in the central complex (CC) of the brain (Homberg, 2015). These cells integrate timing information from bilateral antennae (Guerra et al., 2012) with shifting skylight cues from compound eyes (Heinze and Reppert, 2011, 2012; Heinze et al., 2013). It is thus likely that the output signal from CC is transmitted to the motor system to control flight in the right direction.

5.2 Regulation of sensory systems 5.2.1 Visual system The circadian clock also regulates sensory systems. The visual system shows circadian rhythms in morphology and physiology. The retinal receptor cells and pigment cells in the compound eye often show daily and circadian migration of screening pigments to adapt to day/night light environment. For example, in the bug Triatoma infestans, during the subjective day under constant darkness, screening pigments in photoreceptors and the primary pigment cells form a narrow pupil above the distal tip of the central rhabdomere, while at night, the pupil widens and the rhabdom locates just below the crystalline cone (Reisenman et al., 2002). Thus, a circadian clock apparently regulates both the migration of the pigments forming the “pupil” and the movements of the rhabdom. Similar circadian structural changes in the retina have been reported for the moth Manduca sexta, the cockroach L. maderae, and the cricket G. bimaculatus (Bennett, 1983; Ferrell and Reitcheck, 1993; Sakura et al., 2003). The rhabdom size is also known to show daily changes in some insects (Horridge et al., 1981; Sakura et al., 2003). It is greater during the night and the rhythm persists in constant darkness in the cricket G. bimaculatus (Sakura et al., 2003). The structural changes are also reported for Drosophila optic neuropil. The lamina monopolar cells, L1 and L2, and glia show daily structural changes in both their diameter and synaptic structures, which persist in constant darkness (Pyza and Meinertzhagen, 1995, 1999). The most pronounced circadian changes are seen in the size of L2 axons. The L2 dendrites are longest at the beginning of the subjective day under constant darkness, when the daytime tetrad presynaptic sites are most numerous and L2 axons are swollen (Weber et al., 2009). The control by the circadian clock is partly mediated by PDF, ITP that is produced in the 5th sLNv projecting to the lamina,

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and the catalytic α subunit of the Na+/K+-ATPase that is expressed in lamina glia cells (Damulewicz and Pyza, 2011; Damulewicz et al., 2013). The retinal sensitivity can be monitored by recording an electroretinogram (ERG). ERG shows daily and circadian changes in crickets, cockroaches, and beetles (Fleissner, 1982; Tomioka and Chiba, 1982; Wills et al., 1985). The amplitude of ERG increases at night, suggesting that the sensitivity is higher during this period. The optic tract severance does not eliminate the rhythm (Tomioka and Chiba, 1982), but severance of the optic nerve or destruction in the optic lobe abolishes the rhythm (Fleissner, 1982; Wills et al., 1985). Thus, the rhythm is apparently controlled by the clock in the optic lobe. Visual interneurons in the optic lobe also show circadian rhythms in their responsiveness to light. The light-induced responses of neuronal activity recorded from the distal cut end of the optic stalk, which connects the medulla and the lobula neuropils, show clear circadian changes in the cricket G. bimaculatus (Tomioka et al., 1994). The circadian regulation has been studied in detail by single unit recording in the medulla bilateral neurons that directly connect the right and left medullae of the cricket’s optic lobe (Yukizane et al., 2002). These neurons show a clear circadian rhythm with increased light responsiveness during the subjective night. As previously stated, the circadian regulatory mechanism most likely involves serotonin and PDF. Serotonin decreases neurons’ sensitivity, while PDF increases it, suggesting that serotonin sets the day state and PDF does the night one (Saifullah and Tomioka, 2003a, 2003b). Importantly, the serotonin content in the optic lobe increases during the night, suggesting its release during the day time (Tomioka et al., 1993). 5.2.2 Olfactory system Insect olfactory receptors are located in the antenna, and their circadian rhythm has been studied in Drosophila and other insects. By recording the electroantennogram, sensitivity in Drosophila antennal sensilla was shown to be rhythmically adjustable in a circadian manner (Krishnan et al., 1999). The rhythm is observed in wild-type flies during light-dark cycles and in constant darkness, but is abolished in per01 and tim01 mutant flies, which lack rhythms of adult emergence and locomotor behaviour (Krishnan et al., 1999). The olfactory rhythm is regulated by the clock in the antenna, since the rhythm is eliminated by targeted genetic clock disruption in antennal neurons and targeted rescue of antennal neuron

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oscillators in cyc01 flies through wild-type CYC. This shows that these neurons are also sufficient for olfactory rhythms (Tanoue et al., 2004). The olfactory rhythm is controlled by the rhythmic accumulation of odorant receptors in olfactory sensory neuron dendrites, which is regulated by clock-dependent rhythms in G protein-coupled receptor kinase 2 abundance (Tanoue et al., 2008). Similar antennal olfactory rhythms were reported for mosquitoes, tsetse flies and cockroaches (Rund et al., 2013; Saifullah and Page, 2009; Van der Goes van Naters et al., 1998). In cockroaches, the antennal receptor cells have circadian rhythm, but their synchronization is controlled by the circadian clock in the optic lobe (Page and Koelling, 2003; Saifullah and Page, 2009). As olfaction is essential for food acquisition, social interactions, and predator avoidance in many animals, circadian regulation of olfactory systems could have profound effects on the behaviour of organisms that rely on this sensory modality.

5.3 Regulation of learning and memory Time-dependent memory formation has been known since Beling’s first experiment in which honey bees were trained to visit a feeding place at a specific time of day (Beling, 1929). Her experiment revealed that the bees can memorize the time of feeding. This time-related memory is apparently clock-dependent because when the bees were transferred to a different time zone, they still visited for feeding at the original local time at which they were trained. Recent studies revealed that learning and memory are clock-dependent in the cockroach L. maderae (Decker et al., 2007). Cockroaches can be trained by either classical or operant conditioning in association with olfactory memory tasks. The memory acquisition by classical conditioning occurs in a time-of-day-dependent manner but the acquired memory can be recalled at any time (Decker et al., 2007). In contrast, operant conditioning allowed the cockroaches to learn the task at any circadian phase, but the recall of the acquired memory was linked to the circadian phase when the training was performed (Garren et al., 2013). Optic lobe ablation rescued the deficit in memory acquisition at a time the animals normally cannot learn and rescued the animal’s ability to recall a memory formed by operant conditioning at a phase where memory was not normally expressed (Lubinski and Page, 2016). Thus, the optic lobe clock plays an important role both in memory acquisition and recall.

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5.4 Regulation of photoperiodic time measurement Most insects show seasonal physiological changes in agreement with seasons: diapause is the characteristic state to withstand harsh conditions in winter. The changes are often controlled by photoperiods of which changes are processed by a mechanism called photoperiodic clock (Saunders et al., 2002). It is now widely accepted that the circadian clock is involved in the photoperiodic clock (Saunders, 2012). There are several models which hypothesize the photoperiodic time-measurement based on the circadian clock. Two most well-known are the external coincidence model and the internal coincidence model. The former assumes a particular phase in the circadian clock in which light exposure induces long-day responses, while the latter assumes two circadian clocks synchronizing to dawn and sunset, respectively, and their specific phase relationship induces long-day or short-day responses (Saunders et al., 2002). Although the actual mechanism of photoperiodic time-measurement still remains to be elucidated, the involvement of the clock is already confirmed by molecular and physiological experiments. In the fly Chymomyza costata, malfunction of the circadian clock by mutation of per and tim resulted in a loss of normal photoperiodic responses (Kostal and Shimada, 2001; Pavelka et al., 2003). In the cricket Modicogryllus siamensis, RNAi-mediated gene silencing of per resulted in abnormality of locomotor rhythms and disruption of both long-day and short-day responses at nymphal development (Sakamoto et al., 2009). Similarly, disruption of photoperiodic responses after treatment with clock gene RNAi has been reported in bean bugs (Ikeno et al., 2010, 2011). However, how the photoperiodic clock measures the length of the day or night with the aid of circadian clocks remains to be elucidated.

6. Conclusion and future perspectives: Evolution and diversification of circadian clocks in insects Insects live in various environments to which they have adapted (Whitfield and Purcell, 2012). Circadian clocks may play roles in those temporal adaptations, and the clock itself may have diverged during the course of evolution. At molecular levels, the oscillatory mechanism varies considerably among different phylogenetic groups (Tomioka and Matsumoto, 2015). Although the major frame of the clock consists of clock genes, per, tim, Clk, cyc, and cry in most insects, their roles show variation in higher

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groups. CRY is one of those having diversified roles. Most species have two crys, cry1, and cry2 but, Drosophila possesses only cry1, which encodes a photoreceptor for entrainment in the central clock neurons (Yuan et al., 2007). Hymenopteran species only have cry2, which probably works together with per to form the negative feedback loop because of the lack of the tim gene (Rubin et al., 2006; Zhan et al., 2011). Interestingly, in the monarch butterfly, CRY2 forms a complex with PER to enter the nucleus and represses CLK/CYC-mediated transcription because TIM is rhythmically expressed but stays in the cytoplasm (Zhu et al., 2008). In the cricket, one of the lower groups of insects, cry1 and cry2 have a unique role—they form another negative feedback loop that can function-independent of the per/tim loop (Tokuoka et al., 2017). Another feature of diversification is the role of Clk and cyc. In Drosophila, Clk is rhythmically expressed (Cyran et al., 2003), but in phylogenetically lower groups of insects, including crickets and firebrats, cyc is rhythmically expressed as well (Kamae et al., 2010; Uryu et al., 2013). Considering that Bmal1, a paralogue of cyc, is rhythmically expressed in vertebrates, those lower insects have a clock with an ancestral feature closer to vertebrates. Interestingly, in crickets, when cyc is downregulated by RNAi, Clk starts to oscillate with a period of 24 h (Uryu et al., 2013). Thus, the cricket’s clock may have already started to diverge from the ancestral insect’s clock. Interestingly, the Antarctic midge Belgica antarctica, the only insect endemic to Antarctica, shows daily rhythm with activity during warm phase, but its clock genes, per, tim, Clk, and vri are rhythmically expressed neither in field conditions nor in any photoperiodic conditions (Kobelkova et al., 2015). Thus, this polar species most likely lost the clock oscillatory mechanism. Photic entrainment mechanisms are also an important issue relating to diversification of insect circadian clocks. Lower phylogenetic groups of insects rely only on external photoreceptors for the photic entrainment of the clock (Komada et al., 2015; Page, 1985), while higher groups, such as flies and butterflies, also use CRY1 (Emery et al., 1998; Zhu et al., 2008). Interestingly, in the firebrat Thermobia domestica, a Thysanura species, the cry1 gene could not be found in its genome (Misof et al., 2014). Thus, the entrainment through external photoreceptors seems more primitive. Recently, the compound eye-dependent reset of the clock has been shown to be achieved through c-fos up-regulation (Kutaragi et al., 2018), which is reminiscent of light resetting of the vertebrate circadian clock in the suprachiasmatic nucleus, although the process of the diversification of circadian photoreceptors is an open question.

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However, data on the diversification of insect clocks are still fragmented. To understand the diversification of insect circadian clocks during the course of evolution, comparison of molecular and cellular clock mechanisms should be performed across various taxonomic insect groups. The recent development of molecular techniques, such as RNA-seq using the next generation sequencer, and genome editing technology, including ZFN, TALEN, and CRISPR/Cas9, is now gradually promoting molecular studies using nonmodel insect species. The results of these studies will provide information on the evolution and diversification of the clock system in insects. One important question is the phase of circadian oscillation of clock genes. In insects, per, tim, cry2 generally show a peak during the subjective night while cyc or Clk during the subjective day (Tomioka and Matsumoto, 2015). In contrast, vertebrates show their clock genes to peak with an opposite phase as their insect counterparts (Isojima et al., 2003). Since insects and vertebrates share homologous genes that are involved in the clock machinery, and their clock mechanisms may have the same ancestral origin. How and when they became oppositely phased are important issues that should be addressed in future studies.

Acknowledgements This work is supported by grants from JSPS to K.T. (JP18H02480) and A.M. (JP16K07444). We thank Taishi Yoshii and Taichi Q Itoh for critically reading an earlier version of the manuscript.

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