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date: 03 August 2020

Evolution and Design of Invertebrate Circadian Clocks

Abstract and Keywords

Invertebrates are an incredibly diverse group of animals that come in all shapes and sizes, and live in a wide range of habitats. In order for all these organisms to perform optimally, they need to organize their daily activities and physiology around the perpetuating day-night cycles that exist on Earth. The circadian clock is the endogenous timing system that enables organisms to anticipate daily environmental cycles and governs these roughly 24-hour cellular and overt rhythms. Given its importance to organismal performance and coordination with external environment, it is not surprising that the circadian clock is believed to be ubiquitous in invertebrates. This chapter will discuss the evolution and molecular designs of the invertebrate circadian clocks and describe our current understanding of the circadian clock neuronal network responsible for interpreting external temporal cues and coordinating cellular and physiological rhythms.

Keywords: circadian clock, insect, crustacean, evolution, molecular oscillator, neuronal network

The rotation of the Earth on its own axis creates a predictable 24-hour cycle of environmental variations that serves as a strong selective force to favor organisms that have evolved an endogenous circadian clock to anticipate these daily cycles. Most scleractinian corals, for example, Acropora millespora, extend their tentacles only during night time to actively feed on drifting prey (Sweeney, 1976; Brady et al., 2011). The retraction of tentacles is not simply a light-inhibited response, as extension and retraction of tentacles shows daily 24-hour rhythmicity even in constant darkness, strongly indicating that it is a clock-regulated behavior. The freshwater crustacean Daphnia pulex relies on the circadian clock to control daily rhythms of vertical migration behavior along the water column for resource tracking and predator avoidance ( Tilden et al., 2011; Rund et al., 2016). The Malaria mosquito vector Anopheles gambiae displays circadian rhythms in antennal olfactory sensitivity to host odorants, which directly correlates with times of blood feeding (Rund et al., 2013a; Maliti et al., 2016). These roughly 24-hour cycles of physiology and behavior are so engrained that even Drosophila melanogaster that have been kept in the dark in the laboratory for 1,300 generations still show robust daily rhythms of locomotor activity upon entrainment in light-dark cycles (Imafuku & Haramura, 2011). In fact, there has been experimental evidence supporting that circadian clocks provide organisms with fitness advantage in a competitive environment (Woelfle et al., 2004; Dodd et al., 2005).

This chapter will not be all encompassing with regard to covering all studies related to invertebrate circadian clocks because the scope is simply too expansive. Our goal is to summarize the current understanding of the evolution and molecular design of the circadian clock, as well as to describe the cellular basis for clockwork using selected invertebrate examples.

(p. 596) The Evolution of the Circadian Clock

Although circadian clocks are believed to exist and have been studied in organisms from all kingdoms of life, from bacteria to higher eukaryotes, the path of its evolution remains an ongoing discussion. Multiple hypotheses were proposed to explain why the circadian system might have evolved. Perhaps the most discussed hypothesis is that the circadian clock restricts DNA replication and cell division to night-time in order to reduce probability of UV-induced DNA damage and genome mutation (Ozgur & Sancar, 2003; Oztürk et al., 2009; Papp et al., 2015). This hypothesis is centered on the fact that cryptochrome, a blue-light-sensitive flavoprotein involved in circadian light entrainment in plants as well as in some animal lineages (Somers et al., 1998; Emery et al., 1998; Yuan et al., 2007; Zhu et al., 2008), is postulated to have evolved from prokaryotic photolyases (Oztürk et al., 2007; Zoltowski et al., 2011; Czarna et al., 2013; Kavakli et al., 2016). These are light-activated enzymes that can catalyze the repair of DNA damage. In some vertebrate and invertebrate lineages, gene duplication and functional divergence have led to two types of CRYPTOCHROME (CRY) proteins: one of which remains a light-sensitive protein that regulates firing rate of clock and arousal neurons (Fogle et al., 2011; Baik et al., 2017) and mediates light-activated degradation of circadian transcription factors to enable clock resetting (Koh et al., 2006; Peschel et al., 2006, 2009). Interestingly, although the other type of CRY proteins no longer appears to be light sensitive and has evolved to function as transcriptional repressor of circadian clock-regulated genes (Zhu et al., 2008; Koike et al., 2012), it has retained the ability to bind and sense UV-damaged DNA (Ozgur & Sancar, 2003; Papp et al., 2015), reflecting its photolyase origin. The evolution of CRY will be discussed further later.

A second hypothesis postulates that the circadian clock, together with metabolic enzymes that remove reactive oxygen species (ROS), evolved as a result of the Great Oxidation Event (GOE), which took place approximately 2.33 billions ago (Wang et al., 2011; Kim et al., 2012; Edgar et al., 2012; Wu & Reddy, 2014; Luo et al., 2016). Others have estimated that significant oxygenation of the Earth’s surface happened 600 million years prior to the GOE (Crowe et al., 2013). Presumably, atmospheric oxygenation occurred as a direct consequence of the evolution of oxygenic photosynthesis in bacteria (Dismukes et al., 2001; Buick, 2008; Hohmann-Marriott & Blankenship, 2011). The GOE would have led to a significant decline in and expansion of anaerobic and aerobic life forms, respectively. Assuming that the cycles of cellular production and consumption of oxygen and the fluctuating levels of dissolved oxygen in aquatic environment are closely tied to day-night cycles (Bamforth, 1962; Schilling & Jacobson, 2015), it is likely there was positive selection for a circadian timing mechanism to anticipate and respond to day-night cycles, thereby responding to oxidative stress and maintaining redox homeostasis in cellular environments (Edgar et al., 2012; Wu & Reddy, 2014; Diamond et al., 2017). In support of the hypothesis that maintaining daily redox rhythm and homeostasis was the key selective force for the evolution of the circadian timing system, Edgar et al. (2012) discovered that the oxidation states of the highly conserved peroxiredoxin (PRX) proteins, which are antioxidants important for regulating intracellular hydrogen peroxide levels, display circadian oscillations in every organism found to possess a circadian clock. This includes the smallest known alga Ostreococcus tauri, which possesses a clock with some homology to the plant clock (O’Neill et al., 2011). Interestingly, PRX oxidation cycle closely aligns with circadian cycle in O. tauri even in the absence of gene transcription, suggesting that it may represent the ancestral circadian timing system from which more derived circadian timekeeping mechanisms, that is, oscillators relying on transcriptional-translational feedback loops (TTFL), are built on and/or integrate with (Wu & Reddy, 2014). This would explain the curious lack of orthology in characterized circadian clock components in species from different kingdoms, for example, fungi, plants, and animals, which likely evolved independently. The fact that the functions of many key circadian clock proteins have been shown to be regulated by redox status (e.g., Rutter et al., 2001; Froy et al., 2002; Berndt et al., 2007; Asher et al., 2008; Nakahata et al., 2008; Zoltowski et al., 2011; Czarna et al., 2013; Fogle et al., 2015) also supports the assertion that the circadian clocks we observed today have evolved on the platform of redox cycles.

Finally, the third hypothesis posits that the circadian clock evolved due to the requirement for metabolic and cell cycle synchronization between a prokaryote and an archaebacterium when they fuse to give rise to an early eukaryotic ancestor (Reitzel et al., 2013). Evolution of an endogenous timer to provide temporal organization would have certainly increased the fitness of early eukaryotes. (p. 597) However, given that the most favored hypothesis with regard to the origin of eukaryotes is that they evolved from a lineage of unusually complex archaea (Forterre, 2011; Koonin, 2015; Spang et al., 2015), rather than the fusion of a prokaryote and an archaebacterium, suggests that this scenario is perhaps the least likely.

The Design of the Circadian Timing System and the Molecular Oscillator

The basic architecture of the circadian timing system can be divided into three parts: input, the oscillator, and output. The design and molecular mechanisms underlying the inner workings of the invertebrate oscillator have been under intense investigation since the first characterization of Drosophila melanogaster “clock” mutants by Konopka and Benzer (1971), and subsequent cloning of core “clock” genes (Bargiello et al., 1984; Reddy et al., 1984; Zehring et al., 1984; Sehgal et al., 1994; Myers et al., 1995; Allada et al., 1998; Darlington et al., 1998; Emery et al., 1998; Price et al., 1998; Rutila et al., 1998; Blau & Young, 1999; Cyran et al., 2003; Glossop et al., 2003; Kadener et al., 2007; Lim et al., 2007). Because of the versatile genetic tools available in Drosophila, studies to identify and characterize core clock genes have largely been performed in the Drosophila model. It is only until recently, with the development of high-throughput genomic sequencing (Reuter et al., 2015; Goodwin et al., 2016) and engineering technologies (Gaj et al., 2013) that the characterization of circadian oscillators in other invertebrates are becoming more common. This exciting development will add more taxonomic coverage to our knowledge on circadian clock evolution and physiology. Nevertheless, since the Drosophila oscillator is the most well characterized among invertebrates, we will use it as a model to outline the mechanisms underlying oscillator function and entrainment, and subsequently use a comparative framework to discuss the evolution of the oscillator in invertebrates.

Circadian Oscillator in Drosophila melanogaster

Our understanding of clock mechanisms in invertebrates has been largely based on studies conducted using D. melanogaster as a model system. The oscillator of the Drosophila clock consists of two interlocked TTFLs that produce daily oscillations in clock mRNAs and proteins (Allada & Chung, 2010; Hardin & Panda, 2013; Tataroglu & Emery, 2015) (Fig. 26.1). In the “first” or “feedback” loop, expression of period (per) and timeless (tim), which encode for PAS (Per-Arnt-Sim) domain transcriptional repressors, is activated by a protein complex consisting of two basic helix-loop-helix (bHLH)-PAS transcription factors, CLOCK (CLK) (Allada et al., 1998) and CYCLE (CYC) (Rutila et al., 1998). CLK-CYC heterodimers activate per and tim transcription by binding to E-boxes and other elements in their promoters (Hao et al., 1997; Lee et al., 1998; Lee et al., 1999; Lyons et al., 2000; Darlington et al., 2000; McDonald et al., 2001; Munoz et al., 2002; Paquet et al., 2008). Drosophila per and tim mRNAs begin to accumulate around midday, rise to peak levels in the early night, and reach trough levels during the late night (Allada & Chung, 2010; Hardin & Panda, 2013). A key control mechanism in generating cyclical gene expression that spans 24 hours is the nuclear entry of PER and TIM proteins, which is dependent on DOUBLETIME (DBT; ortholog of mammalian Casein kinase 1δ/ε) and SHAGGY (SGG; ortholog of mammalian GSK3β) kinase activities and the nuclear transport receptor importin α (Martinek et al., 2001; Cyran et al., 2005; Ko et al., 2010; Hara et al., 2011; Jang et al., 2015). This takes place around midnight in most pacemaker neurons. After a controlled delay between the time when per and tim mRNA levels peak, and the time when PER and TIM proteins start to build up in the cytoplasm, PER interacts with TIM, followed by entry into the nucleus, perhaps independently (Meyer et al., 2006), where PER (and perhaps TIM) interacts with and inhibits CLK-CYC activity (Lee et al., 1999; Menet et al., 2010). This repression is relieved upon the eventual degradation of TIM (Koh et al., 2006), and subsequently hyperphosphorylated PER in the nucleus (Ko et al., 2002; Grima et al., 2002), thus initiating another round of CLK-CYC-mediated transcription.

In the “second” or “feed-forward” loop, CLK-CYC activates transcription of two bZIP transcription factors, vrille (vri) and par domain protein 1ε (Pdp1ε) (Cyran et al., 2003; Glossop et al., 2003; Benito et al., 2007). Due to differential kinetics of VRI and PDP1ε protein accumulation, VRI accumulates first and inhibits Clk expression. As VRI level decreases, PDP1ε accumulates and activates Clk transcription, and the cycle of per and tim (and output clock-regulated genes) expression starts again. In addition to the core clock transcription factors mentioned earlier, CLOCKWORK ORANGE (CWO) is another direct CLK target that feedbacks and represses CLK activity by competing (p. 598) with CLK-CYC complexes for E-box binding at circadian promoters (Zhou et al., 2016). Finally, ecdysone-induced protein 75 (E75) and unfulfilled (urf) encode nuclear receptors involved in steroid hormone signaling, which have also been shown to play a role in regulating the circadian oscillator by modulating per and clk expression (Kumar et al., 2014; Jaumouille et al., 2015).

Evolution and Design of Invertebrate Circadian Clocks

Figure 26.1 Schematic representation of the transcriptional translation feedback loop (TTFL) as characterized in Drosophila melanogaster, showing the key components of the molecular oscillator. See text for description.

Although transcriptional control is important for the circadian oscillator to function properly, posttranslational mechanisms, and in particular phase-specific phosphorylation, are critical to the temporal regulation of clock protein levels, their subcellular localization, and transcriptional activities. Posttranslational regulation of clock proteins has proven to be indispensable for extending the duration of the TTFL-based oscillator to approximately 24 hours and for controlling oscillator speed. Through forward and reverse genetic approaches followed by subsequent biochemical analyses, a number of kinases, including DBT (Price et al., 1998; Kloss et al., 1998; Chiu et al., 2008; Kivimäe et al., 2008), Casein kinase 2 (CK2) (Lin et al., 2002; Smith et al., 2008), SGG (Martinek et al., 2001; Ko et al., 2010), and NEMO-like kinase (Chiu et al., 2011; Yu et al., 2011), as well as phosphatases PP1 and PP2A (Sathyanarayanan et al., 2004; Fang et al., 2007) have been identified to modify and regulate clock protein function. In fact, the universal importance of CK1 as a clock component, including in the minimal oscillator of O. tauri, suggests that phosphorylation is likely a primitive clock regulatory mechanism (van Ooijen et al., 2013).

Further complicating the posttranslational regulation of the oscillator, a number of core clock transcription factors (PER and CLK) have recently been shown to be a substrate of O-linked N-Acetylglucosaminylation (O-GlcNAcylation) (Kim et al., 2012; Kaasik et al., 2013). The study of O-GlcNAcylation and its role in cell signaling lags far behind studies on the role of phosphorylation in regulating protein function, but it is now believed that most phospho-proteins are also modified by O-GlcNAc (Hart et al., 2011). Since O-GlcNAcylation also targets serine and threonine residues for modification, as in the case for phosphorylation, it is expected that dynamic interplay between these two modifications will play a role in regulating the function of clock transcription factors and even clock kinases (Wang et al., 2007; Dias et al., 2012). O-GlcNAcylation could provide another route for metabolic input into the oscillator as it is known to be a nutrient sensitive posttranslational modification that is dependent on the influx into the hexosamine biosynthetic pathway (Zeidan & Hart, 2010).

(p. 599) Finally, to add to the molecular mechanisms underlying a robust 24-hour oscillator, a number of ubiquitin ligase components, including SLIMB (ortholog of mammalian β-TrCP), JETLAG, CTRIP, and CULLIN3, were identified to bind to and facilitate the degradation of key clock transcription factors (Grima et al., 2002; Ko et al., 2002; Koh et al., 2006; Lamaze et al., 2011; Grima et al., 2012). Proteasomal degradation of key clock proteins is highly regulated and is often dependent on phase-specific phosphorylation (Chiu et al., 2008). Although not included in this chapter due to limited space, there are other mechanisms that regulate the oscillator function at the posttranscriptional, translational, and chromatin level (reviewed in Bartok et al., 2013; Kwok et al., 2015; Mendoza-Viveros et al., 2016).

Entrainment of the Drosophila Circadian Oscillator

Input pathways enable the circadian timing system to receive external environmental cues, most notably light-dark (Emery et al., 1998; Rieger et al., 2003; Chittka et al., 2013; Yoshii et al., 2015; Arendt et al., 2016; Schlichting et al., 2016; Baik et al., 2017) and temperature (Low et al., 2008; Sehadova et al., 2009; Goda et al., 2014; Chen et al., 2015) cycles, as well as internal metabolic signals (Chaix et al., 2016; Panda, 2016) to entrain the molecular oscillator. The interlocked TTFLs, representing the molecular oscillator in Drosophila, are synchronized to the light-dark cycle through light-induced degradation of TIM (Sehgal et al., 1994), which is dependent on interaction with the photoreceptor type CRY, herein denoted as CRY(P) (Ceriani et al., 1999; Emery et al., 2000; Busza et al., 2004; Peschel et al., 2006; Peschel et al., 2009). Degradation of TIM was hypothesized to depend on tyrosine phosphorylation (Sehgal et al., 1994), although the exact tyrosine target site has not been identified. In addition to CRY(P)-dependent light input, entrainment of the circadian clock neuronal network (CCNN) also relies on the integration of other light inputs from opsin-based photoreceptors in the visual system: the compound eyes, ocelli, and the Hofbauer-Buchner (HB) eyelets (Rieger et al., 2003; Schlichting et al., 2016). The manner in which the signals from different photoreceptors and parts of the visual system are integrated to entrain the circadian clock will likely vary under different conditions, for example, time of day and seasons (Kronfeld-Schor et al., 2013; Schlichting et al., 2016).

It has been shown that zeitgebers, for example, temperature and light-dark cycles, will interact to influence entrainment (e.g., Gentile et al., 2013), but future investigations are necessary to examine interactions between different zeitgebers at the neuronal level (e.g., Zhang et al., 2010) to shape clock output and when these interactions are ecologically relevant. Input elements allow the endogenous oscillator, which can free-run independent of input pathways, to synchronize with the external and internal metabolic environments and orchestrate cellular rhythms in physiology. More important, input pathways provide the necessary plasticity to the circadian timing system to allow for resetting of the oscillator when organisms migrate to a different time zone or encounter changes in internal metabolic conditions.

Molecular Clockwork in Invertebrates

Core oscillator proteins, including PER, TIM, CLK, CYC, and CRY, represent an important protein module in invertebrates that integrate environmental and metabolic signals to regulate cellular physiology, and this module is naturally predicted to be conserved throughout invertebrate lineages and perhaps even throughout metazoan evolution. Surprisingly, comparative analysis of the oscillator protein module from pre-metazoans (choanoflagellates) to various metazoan lineages revealed significant divergence (Fig. 26.2), highlighting unexpected level of plasticity in oscillator design. See legend of Figure 26.2 for methods of comparative phylogenomic analysis. As mentioned earlier, perhaps this is possible since the TTFL-based oscillator was built on the existing platform of daily redox cycles (Edgar et al., 2012) or perhaps even phosphorylation cycles (Nakajima et al., 2005; van Ooijen et al., 2013).


Focusing specifically on core oscillator proteins, our comparative analysis suggests that TIMEOUT (TIMELESS2) appears to be one of the most primitive components of the clock system. TIMEOUT is present even in the genomes of the two choanoflagellates (Salpingoeca and Monosiga), which are single-celled relatives of metazoans. To date, whether TIMEOUT is part of the circadian oscillator in clock models remains a controversy, but there are some evidence supporting its role in circadian entrainment and/or output pathways (Barnes et al., 2003; Ünsal-Kacmaz et al., 2005; Benna et al., 2010). Perhaps proven more conclusively, TIMEOUT has been shown to regulate (p. 600) (p. 601) DNA replication and maintain chromosome stability (Ünsal-Kacmaz et al., 2005; Gotter et al., 2007). This likely represents its ancestral function, especially given that PER, the known interaction partner of TIM, is not present in the genomes of nonbilaterians and hemichordates (Fig. 26.2). It is possible that TIMEOUT collaborates with 6-4 photolyase to regulate light and UV-based DNA damage response in pre-metazoan species, and later evolved in metazoans to serve as a link coordinating circadian timekeeping and DNA replication/repair, perhaps even in collaboration with CRY(R). This would support the hypothesis that avoidance of UV-based DNA damage is a strong selective force for the evolution of circadian clock. New biochemical data on the role of TIMEOUT in either pre-metazoans or early metazoans will reveal the possible role of TIMEOUT in early clocks. So far, expression data in Nematostella vectensis indicates that Nvtimeout does not show apparent circadian cycling (Reitzel et al., 2010).

Evolution and Design of Invertebrate Circadian Clocks

Figure 26.2 Presence of circadian clock genes in the Metazoa. The presence and possible absence of genes in 34 metazoan species and two pre-metazoan choanoflagellate outgroups are determined from the gene family trees generated by OrthologID (Chiu et al., 2006) using the full data set of 1,047,986 gene models from Borowiec et al. (2015) and mapped onto the species tree. Individual gene trees are not shown here but will be provided upon request. Species tree shown is a maximum likelihood (ML) tree estimated using the best-fitting empirical model in RAxML v8.1 (Stamatakis, 2014) based on the “TaxaMin30” subset in Borowiec et al., (2015) with 609 loci and at least 30 taxa present for a total length of 199,667 amino acids. Silhouettes for the different species are public domain images obtained from PhyloPic ( Core clock genes are denoted using the Drosophila melanogaster gene nomenclature. 6-4 represents 6-4 photolyase. cry(p) and cry(r) denote photoreceptor-type cry and transcriptional repressor-type cry respectively. The boxes for 6-4, cry(p) and cry(r) were merged for Amphimedon since, although there are two genes with homology to 6-4 photolyase and cry in the Amphimedon queenslandica genome, they fall outside and is sister to the clade encompassing 6-4 photolyases, cry(p) and cry(r). Although unlikely, absence of genes in our analysis could be a consequence of inaccurate gene models from genome annotation.


Tim is a paralog of timeout, and it was initially identified and characterized in Drosophila (Sehgal et al., 1994). Although originally thought to be insect specific, it is actually present in other bilaterians (Fig. 26.2 and Rubin et al., 2006). Among the genomes of the deuterostomes we analyzed, it is only found in Strongylocentrotus, indicating a widespread loss of tim in that clade. Among the protostomes, loss of tim also appears common in various lineages. In some cases, for example, the Hymenopterans (Apis and Bombus), it is believed that CRY(R) replaces the role of TIM as the partner of PER to inhibit CLK-CYC transcription of circadian genes, similar to the case in mammals (Yuan et al., 2007; Zhu et al., 2008; Sandrelli et al., 2008) (Fig. 26.3). If this is the case for all the species that have lost TIM, we should observe a reciprocal presence of CRY(R) in those lineages, given both TIM and CRY(R) are both present in early bilaterians. But since that was not true in all cases (Fig. 26.2), the presence of a variety of strategies to impose the feedback loop is expected. Furthermore, since TIM plays a role in light entrainment and resetting of the oscillator, the loss of TIM would suggest an alternative strategy for the oscillator to receive light signal, for example, opsin-based phototransduction.

Evolution and Design of Invertebrate Circadian Clocks

Figure 26.3 Schematic diagrams showing three different designs of the molecular oscillator characterized in the monarch butterfly (Lepidoptera, Danaus plexippus) (Yuan et al., 2007; Zhu et al., 2008; Markert et al., 2016) and mosquito (Diptera, genus Anopheles, Aedes, Culex, Wyeomyia) (Gentile et al., 2009; Rund et al., 2013b; Meireles-Filho & Kyriacou, 2013; Tormey et al., 2015), Drosophila melanogaster (Diptera) (Hardin & Panda, 2013; Tataroglu & Emery, 2015; Andreani et al., 2015), as well as honeybee and wasp (Hymenoptera, Apis mellifera and Nasonia vitripennis) (Rubin et al., 2006; Davies & Tauber, 2016). CRY(P) and CRY(R) represent the blue light-sensitive photoreceptor-type CRY and transcriptional repressor-type CRY, respectively. As Hymenoptera (bee and wasp) does not possess TIM and CRY(P), the pathway for light entrainment is expected to be different from other arthropods. The unknown mechanism for light input is denoted by “?.”

(p. 602)


The other key component of the feedback loop, PER, appears to be absent in pre-metazoans and nonbilaterians, and likely first evolved in chordates (Fig. 26.2). Although lost in tunicates (Oikopleura and Ciona), it is found in the genome of lancelet (Branchiostoma), as well as in most of the species analyzed in our phylogenomic analysis. Since the identification of period mutants in Drosophila (Konopka & Benzer, 1971), PER has been shown to play a central role in insect clocks (Allada & Chung, 2010). But with the characterization of clocks in other insect species, it now appears that CRY(R) might have replaced PER as the key repressor of CLK-CYC activity in some lineages. Yuan et al. (2007) indicated that there are at least three basic designs of insect clocks with PER perhaps playing the main repressor role or a supporting role. The ancestral insect oscillator, present in butterflies, moths, mosquitoes, and pea aphids (Acyrthosiphon), as well as oscillators from other protostomes, such as Capitella, Pinctada, and Daphnia, have all the core components of the Drosophila oscillator (Fig. 26.2). In addition, they possess CRY(R) (discussed later), which has transcriptional repressor activity and is orthologous to mammalian CRYs. In the monarch butterfly clock, CRY(R) has been shown to replace PER as the major repressor of CLK-CYC activity (Yuan et al., 2007), similar to the activity of CRYs in mammalian clock (Hardin & Panda, 2013) (Fig. 26.3). It is likely that PER, although present in this more ancestral clock design, is delegated to a supporting role, perhaps in stabilizing CRY(R) or facilitating its nuclear translocation to repress CLK-CYC activity. The Drosophila oscillator, as discussed earlier, represents a derived design where CRY(R) is lost, and its repressor role is replaced by PER. Another derived insect oscillator design described by Yuan et al. (2007) is present in bees and wasps, where PER and CRY(R) are present but both TIM and CRY(P) are lost (Figs. 26.2 and 26.3). As Apis per and cry(r) mRNAs both cycle in LD and DD with a similar phase (Rubin et al., 2006), it is unclear whether PER is the primary repressor in this system. Finally, in addition to the three insect oscillator designs as described by Yuan et al. (2007), a fourth type of insect oscillator can be found in Rhodnius, where both PER and TIM are lost, suggesting that CRY(R) likely functions as the key repressor of CLK-CYC activity (Fig. 26.2).

Nematodes, including C. elegans and Brugia, possess the ortholog of per, also known as lin-42 (Fig. 26.2) (van der Linden et al., 2010; Olmedo et al., 2012). However, lin-42 mRNA does not cycle over a circadian day, but instead correlates with progression of developmental stages. Besides its expression pattern, other strong indicators suggesting that PER in nematodes does not play a role in circadian feedback as in other oscillator types include the lack of CLK-CYC orthologs in nematodes, as well as the fact that cycling lin-42 mRNA level in coordination with larval stages does not depend on the presence of LIN-42 proteins (Jeon et al., 1999). Instead of the TTFL mechanism, the circadian oscillator of C. elegans and more ancestral nematodes may be driven by redox state (Olmedo et al., 2012). The natural history of nematodes, for example, soil-dwelling and parasitic lifestyle, may have led to the need to use redox and metabolic status as their main entrainment signals for rhythmicity.


The evolution of CRY and its role in the oscillators in different species have been of great interest. CRY is believed to have evolved from prokaryotic photolyases (Oztürk et al., 2007; Zoltowski et al., 2011; Czarna et al., 2013) and was first identified in the model plant Arabidopsis thaliana as a photoreceptor for blue and UV-A light (Ahmad & Cashmore, 1993). Since then, CRY proteins have been discovered in microbes as well as in animals (Oztürk et al., 2007). Because of the presence of CRY proteins in plants and animals, CRY proteins most likely evolved prior to the divergence of these two kingdoms. When examining the gene tree for CRY proteins (tree not shown due to size but will be provided upon request), which includes CRY(P), CRY(R), 6-4 photolyase, and CRY-DASH (Selby & Sancar, 2006), the two Amphimedon (Sponge) CRY proteins form its own clade that is sister to the clade encompassing CRY(P), CRY(R), and 6-4 photolyase. As a result, without existing biochemical data, it is unclear what the role of these Amphimedon CRY proteins will have on oscillator function and circadian timing, if any. It will be revealing to determine if Amphimedon CRYs are repressors for CLK, which presumably could act as the activator of circadian transcriptome in Amphimedon. Furthermore, it will also be interesting to examine if the 6-4 photolyase in Mnemiopsis (comb jelly), perhaps the most basal metazoan (Borowiec et al., 2015), plays a role in oscillator function. The Mnemiopsis 6-4 photolyase protein belongs to a clade that is sister to CRY(R), suggesting that it could also potentially act as a repressor-type protein, especially since PER is not present in bilaterians. Investigating the function of these basal metazoan CRY proteins will therefore (p. 603) provide insights into the evolution of CRY proteins with regard to timekeeping.

Nematostella (sea anemone) and Acropora (coral) both have representatives of 6-4 photolyases and CRY(R) (Fig. 26.2). In addition, although not shown in Figure 26.2, these two species also possess CRY proteins that are difficult to classify since they form a clade sister to the monophyletic group encompassing 6-4 photolyases and CRY(R). All these clades then combine to form a monophyletic group that is sister to CRY(P). Reitzel et al. (2010) performed some pioneering experiments to examine light sensitivity and diurnal cycling of the different types of CRY proteins discussed here, and the different classes of CRY proteins showed significant differences in these parameters. Whether any of these CRY proteins play a role in CLK repression will need to be confirmed in future biochemical characterizations.

Similar to the situation of TIM, photoreceptor-type CRY(P) is present in both protostomes and deuterostomes, suggesting CRY(P) evolved prior to the divergence of these two clades. Also, similar to TIM, there has been loss of CRY(P) in various lineages. The selective force for these losses is currently unknown, but it could be related to photoreceptors in the visual system becoming more dominant in these lineages and acting in light entrainment of the oscillator.


These bHLH-PAS transcription activators have long been recognized to act as the main driving force for circadian transcription (Allada et al., 1998; Darlington et al., 1998; Hardin & Panda, 2013). Depending on the species, either CLK or CYC/BMAL1 would bind to E-box elements and activate transcription of clock-regulated genes (Hao et al., 1997; Wang et al., 2013). Both CLK and CYC are present early on in metazoan evolution, but CLK appears to have evolved earlier as it is already present in Mnemiopsis and Amphimedon (Fig. 26.2). Either CLK activates circadian transcription without CYC as a heterodimeric partner, or there is an unknown protein that replaces its function. This pattern seems to be consistent in bilaterian lineages as well; either both CLK and CYC are lost or only CYC is lost, suggesting that CLK is likely the dominant circadian transcriptional activator in those oscillators.


The interaction of this pair of bZIP transcription factors to compete for V/P- or D-box binding and regulate clk transcription was first characterized in the Drosophila oscillator as the feed-forward loop (Cyran et al., 2003; Glossop et al., 2003). Not surprisingly, since these transcription factors are known to be involved in other developmental and physiological pathways (e.g., Lin et al., 1997; Damiola et al., 2000) in addition to circadian timekeeping, they are quite ubiquitous in metazoans (Fig. 26.2). Except in the two most basal metazoans, Mnemiopsis and Amphimedon, as well as the nematode lineages, PDP1 is present in all species analyzed. Its interacting partner in the Drosophila oscillator, VRI, evolved after the nonbilaterians and bilaterians diverged, but it is present in the nematode lineages. Nevertheless, since the nematodes we analyzed do not possess the clk gene, it is perhaps unlikely that VRI is involved in the oscillator mechanism. It still remains possible that nematode VRI is linked to circadian output.


Zhou et al. (2016) recently observed that CWO acts as a repressor of CLK-CYC activity by blocking its access to E-box elements on circadian genes at specific time of the day. Although previously suggested to be an ortholog of DEC1/2 (Bode et al., 2011), which play regulatory roles in mammalian clock (Honma et al., 2002), our phylogenetic analysis revealed that CWO and DEC1/2 are in fact not orthologs (gene tree not shown but available upon request), even though they share a conserved helix-loop-helix DNA binding domain. CWO appears to have evolved after the divergence of protostomes and deuterostomes, and it is only present in some lineages of protostomes and is especially ubiquitous in arthropods (Fig. 26.2).

Organization of the Circadian Clock Neuronal Network and Control of Rhythmic Output

The circadian timing system in animals is hierarchically organized. The central clock or pacemaker, which in many species is now believed to be composed of a coupled network of neurons in the central nervous system (Dissel et al., 2014; Yao & Shafer, 2014; Liang et al., 2016; Schlichting et al., 2016), communicates with peripheral oscillators found in tissues and organs throughout the body to regulate rhythmic gene expression and coordinates physiological processes. Pacemaker cells possess endogenous, cell-autonomous oscillations in clock gene expression in constant conditions, for example, total darkness, which allow them to transduce temporal information to other cells (Strauss & Dircksen, 2010). While central (p. 604) pacemaker cells can be entrained by environmental signals, for example, light-dark cycle, peripheral oscillators are often entrained by other cues, for example, metabolic input and feeding/fasting cycle (Eckel-Mahan & Sassone-Corsi, 2013; Panda, 2016).

Examinations of circadian pacemakers in invertebrates have largely been performed in arthropods, and they have mostly focused on central pacemakers. Details regarding the cellular basis and structural organization of central pacemakers have emerged from studies in insects, primarily flies, butterflies, crickets, and cockroaches, but are also available from a number of crustaceans, for example, the marbled crayfish and intertidal crabs (Farca Luna et al., 2010; Beckwith et al., 2011). The availability of genetic tools in the model insect Drosophila melanogaster, which permit targeted manipulation of clock neuronal electrical properties (Nitabach et al., 2002) as well as in vivo measurements of intact neuronal activity using optical electrophysiology and calcium sensors (Cao et al., 2013; Yao & Shafer, 2014; Liang et al., 2016, enables researchers to assess the circadian relevance, network properties, and function of the pacemaker neurons and has led to significant advance in the dissection of the Drosophila CCNN. In nonmodel organisms and organisms in which cell-type-specific and developmental-stage-specific genetic manipulations remain a challenge, investigations of pacemaker function and network properties have traditionally relied on immunostaining against clock proteins and neuropeptides (e.g., Sauman & Reppert, 1996; Sauman et al., 2005; Grabek & Chabot, 2012), ablation and surgery to remove specific cells or regions of the brain (e.g., Naylor & Williams, 1968; Abe et al., 1997), and transplantation studies to reactivate rhythmicity in animals without rhythm (e.g., Handler & Konopka, 1979; Page, 1982; Reischig & Stengl, 2003). Furthermore, studies examining the endogenous rhythms of dissected tissues in culture have also been used to confirm pacemaker properties (Page, 1981).

In the following sections, studies concerning the organization of the central pacemakers responsible for overt behavioral rhythms in a range of invertebrates are discussed. Mechanisms of clock output, including neuropeptide signaling, will not be discussed in detail in this chapter, but they have been reviewed in Strauss and Dircksen (2010), Taghert and Nitabach (2012), and Shafer and Yao (2014).

Pacemakers in Lancelets

Considered to be basal in the phylum Chordata and as a close invertebrate relative of vertebrates, lancelets have been used extensively as a model to decipher the development and evolution of vertebrates (reviewed in Koop & Holland, 2008). Consequently, various behavioral and developmental aspects of lancelets were comprehensively studied to enable comparative studies (Ruppert, 1997; Wicht & Lacalli, 2005). On the other hand, very little is known regarding its circadian timing system, with scarce observations made regarding the vertical migration of larvae throughout the day (Wickstead & Bone, 1959). Only relatively recently was lancelet demonstrated to exhibit rhythmic burrowing behavior driven by an endogenous clock (Schomerus et al., 2008). Burrowing activity of Branchiostoma lanceolatum oscillates and peaks in the dark phase when entrained to a 12 hour light: 12 hour dark cycle, and it has been observed to oscillate in the absence of light, a hallmark of circadian regulated behavior. Efforts to localize the putative central pacemaker have placed it within a group of cells in the anterior vesicle near the frontal organs in the central nervous system (Schomerus et al., 2008; Wicht et al., 2010). In situ hybridization revealed rhythmic expression of known clock components only in these cells within whole lancelets. These results combined with the homology between the anterior vesicle of lancelets and the hypothalamic region of vertebrate that contains the master pacemaker indicate a high likelihood that the pacemaker is indeed located inside the anterior vesicle (Lacalli, 1996; Wicht & Lacalli, 2005; Rosenwasser & Turek, 2015). Future experiments are necessary to provide support to this view.

Pacemakers in Crustaceans

Many crustaceans are known to display rhythmicity in various behavioral and physiological processes (reviewed in Strauss & Dircksen, 2010). Studies of the crustacean clock have been conducted mainly using decapod species, for example, lobsters and crayfish. Commonly examined clock outputs include locomotor activity, neuronal firing of the retina via electroretinogram (ERG), and pigment dispersion within the chromatophores (Ringelberg & Servaas, 1971; Page & Larimer, 1972; Fernandez de Miguel & Aréchiga, 1994; Aréchiga & Rodriguez-Sosa, 1998; Fuentes-Pardo et al., 2003; Miranda-Anaya, 2004; Gaten et al., 2008; Chiesa et al., 2010; Zhang et al., 2013). Consistent with the notion that these rhythmic processes are controlled by (p. 605) endogenous clocks, they remain rhythmic in constant conditions and thus were used as markers in subsequent efforts to locate the central pacemakers.

Since genetic tools for targeted manipulation are limited in crustaceans, progress has been slow toward pinpointing the location and understanding the structural organization of the central pacemaker in crustaceans. Early studies to locate the pacemakers have mostly relied on ablation or lesion of structures predicted to contain the circadian oscillators, coupled with behavioral analysis, that is, locomotor activity rhythms, to assess the functionality of the clock in the lesioned animals. More recent studies are taking advantage of immunostaining using antibodies for key clock proteins, such as PER, TIM, and CLK (Escamilla-Chimal et al., 2010). Ablation experiments in crayfish species in the genus Potamobius and Cambarus placed the central pacemakers within the eyestalks since ablated animals showed arrhythmicity in locomotor activity (Schallek, 1942; Roberts, 1944). In support of this finding, the endogenous rhythm of another freshwater crayfish species, Procambarus clarkia, was similarly lost following eyestalk ablation (Sullivan et al., 2009). Animals that previously exhibited robust locomotor rhythm became arrhythmic in constant condition once the eyestalks were removed. Unfortunately, this notion is not without ambiguity. Eyestalk ablation in P. clarkii was previously observed to not consistently abolish locomotor rhythms (Page & Larimer, 1975). A proportion of the ablated animals retained the endogenous rhythm while others exhibited arrhythmicity. Nevertheless, studies of other decapods, though scarce, have also localized the pacemakers to the eyestalk. For example, eyestalk ablation in the crab genus Gecarcinus and Carcinus abolished the endogenous activity rhythm (Bliss, 1962; Naylor & Williams, 1968).

Perhaps the strongest evidence that the pacemaker resides in the eyestalk came from the lobster, Homarus americanus, in which PER proteins were not detected in the brain but were observed to be cycling in the eyestalk (Grabek & Chabot, 2012). However, whether PER cycling is critical to the pacemaker in H. americanus depends on whether PER, and not CRY(R), is the main repressor of the oscillator in this species. In the crayfish P. clarkii, however, PER, TIM, and CLK were all observed to be present and cycling in both the eyestalk and the brain (Escamilla-Chimal et al., 2010), suggesting that some crustaceans likely do not contain one central pacemaker responsible for locomotor activity rhythm, but rather, multiple coupling oscillators reside in various tissues cooperate to produce the overt locomotor rhythm. In support of this, severing neural connection surrounding the supraesophageal ganglion abolished locomotor rhythm in P. clarkii (Page & Larimer, 1975). Given the existing data, the eyestalk remains the most likely location for the main pacemaker in some crustaceans, similar to the pigment dispersing factor (PDF)-positive lateral neurons in Drosophila melanogaster (Helfrich-Förster & Homberg, 1993).

A second, well-documented overt rhythm driven by the endogenous clock in decapods is the amplitude of ERG, which reflects the rhythmic migration of the retinal pigments (Aréchiga & Fuentes, 1970; Fanjul-Moles & Prieto-Sagredo, 2003). Efforts to locate the pacemaker underlying this rhythm, which could be coupled to other pacemakers that regulate locomotor activity rhythms, have identified multiple oscillatory centers crucial for proper generation of ERG rhythm. First, removal of the supraesophageal ganglion abolished ERG rhythm (Aréchiga et al., 1973). Similar arrhythmicity was observed in lesions of the central part of the optic tract (Larimer & Smith, 1980). On the other hand, isolated eyestalk exhibited and maintained ERG rhythm in both entrained and constant conditions when kept in media (Aréchiga & Rodriguez-Sosa, 1998). Interestingly, the same authors observed in a previous experiment that when transplanted, the grafted eye adopted the host’s rhythm. Likely, crustaceans utilize a multicomponent, coupled-pacemaker system to generate and sustain the endogenous ERG and other physiological rhythms.

Pacemakers in Insects

Arguably, the location and neuronal organization of the circadian pacemaker are best characterized in insects. Extensive data have been obtained for Orthoptera (crickets), Blattodea (cockroaches), Lepidoptera (butterflies and moths), and especially Diptera (flies) (Numata et al., 2015). As characterized in Drosophila melanogaster, these organisms utilized a coupled network of clock neuronal groups that display hierarchical importance to drive overt rhythms. Neuroanatomical analyses of different insect groups as well as comparative analyses have been reviewed in Tomioka and Abdelsalam (2004); Helfrich-Förster (2004, 2005); Sandrelli et al. (2008); Heinz and Reppert (2012); Heinz et al. (2013); Tomioka (2014); Numata et al. (2015); Yoshii et al. (2015); Stengl and Arendt (2016); and Reppert et al. (2016).

(p. 606) In cockroaches and crickets, key elements of the master pacemaker are believed to be located in the optic lobes. In the cockroach Rhyparobia maderae, removal of the optic lobes consistently abolished locomotor activity rhythm (Nishiitsutsuji-Uwo & Pittendrigh, 1968). Similar results have been obtained with Teleogryllus commodus and Gryllus bimaculatus, two cricket species that are commonly used for circadian research (Sokolove & Loher, 1975; Tomioka & Chiba, 1984). Removal of a single optic lobe, however, does not generate arrhythmia in either group of organism, indicating that each optic lobe contains a fully functioning master oscillator (Nishiitsutsuji-Uwo & Pittendrigh, 1968; Page, 1978; Okamoto et al., 2001). Cementing the optic lobe as the locus of the clock, optic lobe-less and arrhythmic animals were able to regain normal locomotor rhythm following optic lobe transplantation surgeries (Tomioka, 2014).

Structurally, the optic lobe consists of three distinct neuropils: the lamina, medulla, and lobulla. In cockroaches, only lesions within the medulla and lobula complex generated arrhythmia, while lesions within the lamina did not affect normal rhythm (Roberts, 1973). Subsequent transplantation and comparative studies identified a small section between the medulla and the lobulla, the accessory medulla (AME), as the site containing key elements of the master oscillator (Reischig & Stengl, 2003; Homberg et al., 2003; Helfrich-Förster, 2004). Transplanting the AME into cockroaches lacking the optic lobes restored the locomotor activity in these animals (Reischig & Stengl, 2003). The role of the AME as the master oscillator in cockroach has prompted detailed analysis of the neuronal organization of this structure. The investigations into the specific circadian function for different groups of neurons within the AME have been reviewed in Stengl et al. (2015). In crickets, the lamina and medulla, excluding the AME, appeared to be the locus of the master oscillator as the removal of these regions caused arrhythmia in various species (Tomioka & Chiba, 1984; Abe et al., 1997; Okamoto et al., 2001). Whether the lamina or the medulla individually contains the central pacemaker has yet to be determined.

In lepidopterans, the central pacemaker has been localized to the midbrain. Early on, ablation studies using silkmoths recognized the importance of the midbrain in generating overt behavioral rhythms, such as flight activity and eclosion (Truman & Riddiford, 1970; Truman, 1972, 1974). Following these initial studies, expression patterns of known clock components were used as key evidence to identify four neurons within the pars lateralis in the moth Antheraea pernyi as key elements of the central pacemaker (Sauman & Reppert, 1996). Supporting evidence was later obtained in the monarch butterfly, Danaus plexippus, where the same neurons were also identified as clock neurons (Sauman et al., 2005; Zhu et al., 2008).

In dipterans, the location and neuronal organization of the central pacemaker have been elucidated in details in Drosophila melanogaster. Early transplantation and mosaic experiments showed that the master oscillator driving the bimodal locomotor activity rhythm of D. melanogaster was localized within the brain (Handler & Konopka, 1979; Konopka et al., 1983). Subsequent analysis of mutants lacking the optic lobes indicated that they remained rhythmic, though with altered period length, thus substantiated this view (Helfrich & Engelmann, 1983; Helfrich, 1986). Within the brain, the central pacemaker consists of a coupled network of 150 neurons, subdivided into seven groups according to size and anatomical location (Helfrich-Förster et al., 2007; Tomioka & Matsumoto, 2010; Yao et al., 2012). In brief, there are three clusters of dorsal neurons: DN1 (16 cells), DN2 (2 cells), and DN3 (40 cells of varying sizes). The four clusters of lateral neurons (LNs) are further divided into large and small ventrally located LNs (l-LNvs [4 cells], s-LNvs [5 cells]), dorsally located LNs (LNds [6 cells]), and posteriorly located LNs (LPNs [3 cells]).

The CCNN receives light entrainment signals through photoreceptors expressed in the visual system (compound eyes, ocelli, and Hofbauer-Buchner [HB] eyelet) and the extraocular photoreceptor CRY(P), which are expressed in a subset of clock neurons, including LNvs, and LNds. s-LNvs in particular have been shown to function as the master regulator of the CCNN (Grima et al., 2004; Stoleru et al., 2004). Flies in which the oscillators are disrupted in these neurons fail to maintain robust rhythms in constant conditions, indicating that these neurons are necessary to drive endogenous locomotor activity rhythms (Renn et al., 1999; Nitabach et al., 2002). s-LNvs drive the morning peak of locomotor activity and synchronize the CCNN, including the cells (LNd and fifth s-LNv) controlling the evening peak of activity, by releasing the neuropeptide PDF (Lear et al., 2005; Hyun et al., 2005; Rieger et al., 2006; Schlichting et al., 2016).

Recent studies highlighted the complex neuronal coupling that underlies the proper functioning (p. 607) and plasticity of the CCNN. By manipulating oscillator speed of clock neurons other than the s-LNvs using genetic approaches, researchers were able to modulate the locomotor activity rhythm in transgenic flies (Yao & Shafer, 2014; Dissel et al., 2014, Guo et al., 2014; Schlichting et al., 2016). The current model therefore postulates that the behavioral output results from interplay between independent clusters of pacemaker neurons rather than being dependent on one dominant cluster. Finally, the CCNN projects to and communicates with other brain regions, including the DH44-expressing cells in the pars intercerebralis (PI) and cells expressing Leucokinin neuropeptide (LK) and its receptor (LK-R), for example, in the lateral horn, to impose temporal control on physiological and behavioral output processes (Cavanaugh et al., 2014; Cavey et al., 2016).


High-throughput sequencing technologies are rapidly enabling the identification of clock and photoreceptor genes important for regulating the circadian oscillator (e.g., Srivastava et al., 2010; Ryan et al., 2013) and uncovering output molecular pathways that are regulating rhythmic physiology and behavior (e.g., Hughes et al., 2012; Rodriguez et al., 2013; Rund et al., 2013b, Rund et al., 2016) in model and nonmodel organisms alike. This creates opportunities to generate new hypotheses to examine the evolution as well as the molecular and cellular basis of circadian clocks in a comparative framework. Detailed analysis of the CCNN and biochemical characterizations of identified clock components to examine their functional roles and interactions with other cellular machineries will continue to promote our understanding of the complex circadian control of cellular rhythms. Now that we have a basic understanding of the circadian timekeeping system, new challenges will be to investigate the mechanisms by which the circadian clock can adapt to function effectively in a latitudinal cline (Sandrelli et al., 2007; Tauber et al., 2007; Svetec et al., 2015; Rivas et al., 2016) and manifest plasticity in response to natural or even human-made environmental variations, for example, seasonal cycles and global climate change.


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