CIRCADIAN REGULATION OF METABOLISM

Clock Control of Carbohydrate Metabolism

Dramatic time-of-day-dependent oscillations in glucose metabolism are observed in both humans and rodent models, at both the whole body and organ/cellular levels. During periods of increased physical activity, non-insulin mediated glucose utilization increases; the relative contribution of aerobic to anaerobic utilization being dependent upon exercise intensity (Alberts et al. 2006; Calvo, et al. 2008; Rose and Richter 2005). Similarly, food consumption results in insulin-mediated glucose disposal (Woerle, et al. 2003). It is therefore not surprising that indirect calorimetry reveals increased glucose utilization during the awake period (relative to the sleep period). However, as highlighted in the preceding section, recent studies strongly support the concept that rhythms in glucose metabolism are not mediated purely by fluctuations in behaviors. Additional evidence includes the observation that circulating glucose levels increase prior to waking, in both humans and rodents, an event known as the dawn phenomenon (Bolli, et al. 1984). Similarly, rhythms in blood glucose levels persist when rats are fasted (La Fleur, et al. 1999). Collectively, these observations reveal an important role of an endogenous circadian mechanism in whole body glucose metabolism. Indeed, surgical ablation of the central clock (SCN) or genetic manipulation of circadian clock components disrupts glucose homeostasis (La Fleur et al. 1999; la Fleur, et al. 2001). In the case of the dawn phenomenon, multiple studies suggest that the SCN-paraventrciular nucleus-autonomic nervous system axis plays a critical role in daily rhythms in hepatic glucose output, as recently reviewed by Kalsbeek et al (Kalsbeek, et al. 2014).

Glucose homeostasis is achieved through the coordinated regulation of exogenous (ingestion/digestion/absorption) and endogenous (gluconeogenesis) glucose input versus disposal (utilization) mechanisms. Evidence exists that the hepatocyte circadian clock plays an important role in several processes involved in glucose homeostasis, including gluconeogenesis and glycogen turnover. In the latter case, hepatic glycogen levels display a diurnal variation in a variety of species including rats, mice, rabbits, guinea pigs, chickens, and human as highlighted in (Sollberger 1964). Interestingly, time-of-day-dependent rhythms in glycogen levels persist in fasted rodents (albeit at lower amplitude), suggesting that these rhythms are not simply secondary to feeding/fasting cycles (Ishikawa and Shimazu 1976). Not only do glycogen levels oscillate over the course of the day, but so do the activities of key glycogen metabolism enzymes; glycogen synthase displays maximum levels during the dark (active) phase in rodents, whereas glycogen phosphorylase peaks towards the end of the light (sleep) phase (Ishikawa and Shimazu 1980; Peret, et al. 1976). Evidence in support of circadian clock involvement in mediation of these oscillations comes from recent studies by Doi et al, which report diminished oscillations in both hepatic glycogen levels and glycogen synthase expression/activity in Clock^Δ19 mutant mice (Doi, et al. 2010). Similar to glycogen turnover, glucogenogenesis exhibits a diurnal variation; increased rates are observed at the sleep-to-wake phase transition (in comparison to the wake-to-sleep transition), which is paralleled by a diurnal rhythm of phosphoenolpyruvate carboxykinase (PEPCK) activity, a key regulatory enzyme in gluconeogenesis (Kida, et al. 1980). Using hepatocyte-specific BMAL1 knockout mice, Lamia et al found that the hepatocyte circadian clock was critical for rhythms in PEPCK expression (Lamia, et al. 2008). More recently, Zhang et al reported that the core clock component CRY modulates hepatic gluconeogenesis in a time-of-day-dependent manner through regulation of β-adrenergic signaling and cAMP response element-binding protein (CREB) activation (Zhang, et al. 2010).

Both insulin-dependent and insulin-independent whole body glucose disposal exhibits diurnal variations in humans and rodents (la Fleur et al. 2001; Lee, et al. 1992; Whichelow, et al. 1974). Extra-hepatic tissues, such as muscle, contribute in a significant manner to glucose homeostasis. Diurnal variations in both skeletal and cardiac muscle glucose utilization persist ex vivo, suggesting that an intrinsic mechanism may contribute towards these rhythms (Leighton, et al. 1988; Young, et al. 2001a). In the case of the heart, genetic disruption of the cardiomyocyte circadian clock abolishes time-of-day-dependent rhythms in cardiac glucose oxidation, glycolysis, and glycogen synthesis, as well as protein O-GlcNAcylation (an indirect marker of flux through the hexosamine biosynthetic pathway) (Durgan et al. 2011). More recently, Dyar et al reported that decreased soleus and diaphragm muscle glucose uptake and oxidation in skeletal myocyte-specific BMAL1 null mice, concomitant with dysregulation of pyruvate dehydrogenase (Dyar, et al. 2014). Collectively, these observations suggest that circadian clocks within cardiac and skeletal myocytes significantly influence glucose utilization.

Clock-mediated regulation of glucose homeostasis also involves the temporal regulation of endocrine factor release and sensitivity. Insulin and glucagon play important roles in maintaining blood glucose homeostasis (Voet 2004). In vitro studies by Peschke et al. showed that insulin secretion from isolated pancreatic rat islets display a circadian rhythm that originates within the islet (Peschke and Peschke 1998). Further, plasma insulin concentrations in rats exhibit daily oscillations with distinct increments at every meal (Kalsbeek and Strubbe 1998). Studies also show that disruption of the circadian clock causes impaired insulin secretion and consequent hypoinsulinemia (Coomans, et al. 2013; Marcheva, et al. 2010). Using melatonin receptor null mice as a model of circadian clock dysregulation, Muhlbauer et al reported altered circadian rhythms of insulin transcripts and plasma insulin levels (Muhlbauer, et al. 2009). Insulin sensitivity also exhibits a time-of-day-dependence, which appears to be circadian clock dependent. For example, both glucose/insulin tolerance are impaired in SCN ablated rats, as well as Clock^Δ19 mutant and BMAL1 null mice (la Fleur et al. 2001; Rudic, et al. 2004). Similarly, insulin signaling is impaired in various tissues isolated from germline BMAL1 null, PER2 mutant, and cardiomyocyte-specific Clock^Δ19 mutant mice (Anea, et al. 2009; Carvas, et al. 2012; Durgan, et al. 2010). In addition to insulin, glucagon also plays a crucial role in regulating blood glucose homeostasis, and has similarly been shown to display diurnal patterns of release (Gagliardino, et al. 1978; Tasaka, et al. 1980). A study by Ruiter et al demonstrated that 24 hour plasma glucagon concentrations are regulated by feeding and the circadian clock (Ruiter, et al. 2003). Experiments by Bahr et al suggest that melatonin (an important synchronizer of circadian rhythms) influences pancreatic glucagon expression, as well as peripheral glucagon action (Bahr, et al. 2011). Collectively, these observations raise the possibility that cell autonomous circadian clocks contribute towards rhythms in glucose homeostasis, through regulation of endocrine factors (e.g., insulin and glucagon) release and/or sensitivity.

Clock Control of Lipid Metabolism

Similar to glucose, it is well documented that lipid metabolism displays time-of-day dependent rhythms, which align with daily rhythms in behaviors, such as sleep/wake and feeding/fasting cycles. Significant evidence collected using genetic mouse models of clock disruption support the concept that rhythms in lipid metabolism are mediated by intrinsic molecular clocks. The purpose of this subsection is to provide a brief overview of current knowledge regarding clock control of lipid digestion/absorption, as well as oxidative and non-oxidative metabolism of lipids and fatty acids.

Uptake of dietary fat by the body involves mechanical emulsification, lipolytic breakdown, and absorption into intestinal epithelial cells. The digestive tract displays daily rhythms at multiple levels, including gastrointestinal motility, exocrine secretion, macronutrient absorption, and digestive enzyme activities. Using an in situ jejunal loop preparation and isolated primary enterocytes, Pan et al reported higher rates of cholesterol and lipid absorption during periods of high activity (dark phase) in mice and lower absorption during periods of low activity (light phase) (Pan and Hussain 2009). Importantly, this study revealed that Clock^Δ19 mutant mice absorbed similar amounts of macronutrients during both the day and night, indicating a loss of rhythm in intestinal absorption, and supporting a role for the circadian clock in regulating nutrient absorption. Indeed, robust expression of clock genes in different regions and cell types in the gastrointestinal tract have been observed by several groups (Hoogerwerf, et al. 2007; Pardini, et al. 2005; Sladek, et al. 2007). Further, studies have shown that many of the genes involved in lipid uptake and metabolism in the intestine including apolipoprotein B (apoB), apolipoprotein AIV (apo AIV), intestinal microsomal triglyceride transport protein (MTP), and intestinal fatty acid binding protein (FABP) display diurnal rhythms (Pan and Hussain 2007, 2009; Pan, et al. 2010). Additional evidence for clock control of lipid digestion/absorption comes from observations that mice with genetic ablation of Nocturnin (a known clock controlled gene) are resistant to high fat diet induced obesity, due to aberrant chylomicron excretion from intestinal epithelial cells (Douris, et al. 2011). While the importance of the intrinsic clock in regulating lipid metabolism-related functions in the intestinal tract are established, rhythms in these processes and others can be phase-shifted in response to restricted feeding regimens and attenuated by different lighting conditions (e.g., constant darkness or constant light), pointing to the importance of both feeding behavior and the light-dark cycle in time-of-day-dependent regulation of intestinal function (Malloy, et al. 2012; Pan and Hussain 2009; Polidarova, et al. 2011). Moreover, Mukherji et al recently showed that proper functioning of the circadian clock in intestinal epithelial cells is necessary for maintaining homeostasis between the intestinal epithelium and gut microbiota (Mukherji, et al. 2013). The importance of this relationship is showcased by work demonstrating that disrupting the intestinal circadian clock causes gut leakiness and hepatic inflammation (Swanson, et al. 2011). Thus, continued investigation and improved understanding of the intestinal circadian clock is important especially with the newly recognized role of the intestine-microbiome relationship in influencing metabolic syndrome (Vinje, et al. 2014).

In addition to the processes described above, several lipid metabolism species exhibit daily rhythms. For example, circulating non-esterified fatty acids (NEFA) are known to display diurnal rhythms. Multiple studies have reported higher circulating levels of NEFA in rodents during their inactive phase (Shostak, et al. 2013; Stavinoha, et al. 2004). Similarly, humans have higher levels of NEFA at night due to increased lipolytic activity (Schlierf and Dorow 1973). In support of an underlying circadian regulation of NEFA homeostasis, studies comparing circulating NEFA levels in fasted human volunteers revealed that plasma NEFA levels remained higher in the evening compared to the morning, despite lack of food intake (Carroll and Nestel 1973; Gibson, et al. 1975). Further, a recent study by Dallman et al found that plasma lipid levels remain rhythmic in human subjects maintained in constant dim light, even when food intake and wakefulness were kept constant (i.e., hourly iso-caloric food intake and sleep deprivation) (Dallmann, et al. 2012). Taken together, these studies provide strong evidence that rhythms in circulating lipid species (and presumably lipid metabolism) are not simply secondary to behavioral cycles.

Circadian regulation of triglyceride metabolism is achieved in part through the rhythmic gene expression of several enzymes involved in triglyceride biosynthesis (Adamovich, et al. 2014; Shostak et al. 2013). Lipid biosynthesis is known to be regulated by sterol regulatory element-binding proteins (SREBP), a family of membrane-bound, basic helix-loop-helix leucine zipper (bHLH-LZ) transcription factors (Goldstein, et al. 2006). Emerging studies reveal a transcriptional regulatory link between SREBP and the circadian clock. By the use of a combination of genetic loss- and gain-of-function experiments, Le Martelot and colleagues found that REV-ERBα regulates the daily rhythm in activity and expression of SREBP, as well as SREBP targets (e.g., fatty acid synthase [FAS] and acetyl-CoA carboxylase α [ACCα]), independently of feeding regimen (Le Martelot, et al. 2009). Regulation of lipid uptake, biosynthesis, and breakdown are tightly coupled. Therefore it is not surprising that lipid turnover and fatty acid β-oxidation are also diurnally regulated. Enzymes such as carnitine palmitoyltransferase 1 (CPT1), medium-chain acyl-CoA dehydrogenase (MCAD), adipose triglyceride lipase (ATGL), hormone-sensitive lipase (HSL), and diacylglycerol acyltransferase 2 (DGAT2) have been shown to exhibit circadian oscillations in various tissues at the gene expression level (Alenghat, et al. 2008; Filiano, et al. 2013; Shostak et al. 2013; Tsai, et al. 2010). In support of clock control of these genes, Shostak et al have shown that HSL and ATGL oscillations are absent in white adipose tissue isolated from BMAL1 null mice, while Tsai et al have reported loss of DGAT2 oscillations in hearts of cardiomyocyte-specific Clock^Δ19 mutant mice (Shostak et al. 2013; Tsai et al. 2010). More recently, Bass and colleagues have proposed time-of-day-dependent regulation of hepatic β-oxidation through the circadian clock by protein acetylation (as discussed in greater detail in subsequent sections) (Peek, et al. 2013). Taken together, it is clear that cell autonomous circadian clocks likely regulate lipid metabolism at multiple levels.

Clock Control of Protein and Amino Acid Metabolism

An important example of temporal regulation of biologic processes by cell autonomous circadian clocks is DNA synthesis and repair. DNA repair is increased during the active period, when oxidative stress is high, while DNA synthesis is restricted to the less active/sleep phase (Edery 2000). In doing so, the clock minimizes transmittance of mutations to daughter cells. An analogous form of regulation may occur for protein turnover. Oxidative stress is predicted to cause protein damage primarily during the active phase, while increased protein degradation and autophagy are observed during the less active/sleep phase (presumably as a means to remove damaged proteins/organelles, in anticipation of the subsequent active period). The nature of protein rhythms in protein synthesis appear to be less consistent; skeletal muscle net protein synthesis peaks during the active/awake phase, which is in marked contrast to cardiac muscle, for which protein synthesis peaks during the sleep/inactive phase (Garlick, et al. 1973; Rau and Meyer 1975). Consistent with these metabolic observations, several microarray studies have reported marked diurnal variations in genes known to modulate protein turnover, including ubiquitin ligases, proteasome subunits, and autophagy proteins and markers (Duffield, et al. 2002; Reddy, et al. 2006). Classically, stimulation of protein synthesis (e.g., skeletal muscle) and concomitant repression of protein degradation (e.g., liver) during the active/awake phase has been attributed to increased food intake at that period of time (Garlick et al. 1973). Indeed, circulating insulin levels (a potent anabolic signal) are increased during the active phase (Fulks, et al. 1975; Rannels, et al. 1975). Similarly, circulating amino acid levels are elevated during the awake phase, which act both as a substrate and stimulus for protein synthesis (Wurtman, et al. 1968). Interestingly, diurnal variations in circulating amino acid levels persist both during fasting and consumption of protein-free diet, suggesting that ingestion of dietary protein during the active period is not mediating rhythms in circulating amino acids (or protein synthesis) (Fernstrom, et al. 1979). These observations raise the possibility that oscillations in protein degradation are independent of dietary status (i.e., feeding/fasting cycles), releasing amino acids into the circulation in a time-of-day-dependent manner.

Evidence has begun to accumulate, suggesting a role for circadian clocks in the regulation of protein turnover. Indeed, time-of-day-dependent turnover of circadian clock proteins is essential for the functioning of this mechanism. This is exemplified by the E3 ubiquitin ligases F-Box and leucine-rich repeat protein 3 (FBXL3) and FBXL21, which are essential for turnover of CRY proteins (Yoo, et al. 2013). Similarly, microarray analyses highlight multiple components of the ubiquitin/proteasome system as being circadian clock regulated (Duffield et al. 2002; Reddy et al. 2006). One such component, ubiquitin specific protease 2 (USP2; important for deubiquitination of proteins), is considered a direct CLOCK-BMAL1 target gene that influences hepatic gluconeogenesis (Molusky, et al. 2012). Interestingly, this clock output gene oscillates in a number of additional metabolically active tissues, including skeletal and cardiac muscle (Bray et al. 2008a; McCarthy, et al. 2007). As highlighted above, insulin is a potent anabolic signal, and evidence suggests that cell autonomous circadian clocks influence insulin sensitivity; clock-mediated diurnal rhythms in insulin sensitivity may therefore contribute towards oscillations in protein turnover. Similarly, whether clock-driven oscillations in amino acid metabolism (particularly branched chain amino acids, BCAA) impact protein turnover is an attractive hypothesis. Indeed, microarray analyses suggest regulation of the BCAA alpha ketoacid dehydrogenase by the circadian clock at the transcriptional level (unpublished observations).

Clock Control of NAD⁺ Metabolism

NAD⁺ is traditionally considered as a key source of reducing equivalents for numerous oxidation-reduction reactions being interconverted between its oxidized (NAD⁺) and reduced (NADH) forms. However, NAD⁺ may also function as a cell signaling molecule via roles in PTMs; ADP-ribosylation and NAD⁺-dependent deacetylation. For example, NAD⁺ activates sirtuin 1 (SIRT1) deacetylase thereby altering the acetylation status and presumably the function of cellular proteins (Imai, et al. 2000; Landry, et al. 2000). There are two metabolic pathways that control the levels of NAD⁺, the biosynthesis pathway from tryptophan and the salvage pathway (Imai 2010). Notably, the salvage pathway functions to preserve cellular NAD⁺ levels when NAD⁺ is used as a cofactor for signaling reactions. For example, when NAD⁺ is used as a co-factor for SIRT1 it is degraded to nicotinamide (NAM). In the salvage pathway, NAM is converted to nicotinamide mononucleotide (NMN) by the enzyme nicotinamide phosphoribosyltransferase (NAMPT); the rate-limiting enzyme in the pathway. NMN is then reacted with ATP by the enzyme nicotinamide/nicotinic acid mononucleotide adenylytransferase (NMAT) to regenerate NAD⁺. Importantly, the NAD⁺ salvage pathway operates in a circadian fashion because NAMPT gene expression is regulated by CLOCK-BMAL1 binding (Nakahata, et al. 2009b; Ramsey, et al. 2009). The consequence of this is daily oscillations in NAD⁺ and NAD⁺-dependent reactions (e.g., SIRT1 activity). Importantly, daily rhythms in NAMPT and NAD⁺ are absent in tissues and cells from animal models with a nonfunctional circadian clock mechanism and pharmacological inhibition of NAMPT activity also dampens NAD⁺ oscillations (Nakahata, et al. 2009a; Peek et al. 2013; Ramsey et al. 2009). Together, these results demonstrate that the molecular clock plays an important role in regulating the levels of a key cellular metabolite, NAD⁺, which, in turn, can feedback and regulate clock activity (see section on Acetylation). With this said there are still gaps in this area of NAD⁺ biology that require attention. For example, it is not clear what role the molecular clock plays in regulating different sub-cellular pools (nuclear, cytosolic, and mitochondrial) of NAD⁺. This is an important question as most studies have only looked at total cellular or whole tissue levels of NAD⁺ and we now know that organelle compartmentation and microenvironments are critical aspects in control of metabolism. Furthermore, measurements done using isolated organelles (e.g., mitochondrial and/or nuclear fractions) may be invalid as NAD⁺ content and/or the NAD⁺/NADH ratio will likely be altered during the sub-cellular fractionation procedure. However, investigations with a new mitochondrially-targeted poly(ADP-ribose) polymerase 1 (PARP-1) derivative as a tool for NAD⁺ detection have shed some light into mechanisms involved in mitochondrial NAD⁺ generation (Nikiforov, et al. 2011). It is also important that investigators explicitly report the nutritional state (fed vs. fasting), feeding strategy (ad libitum vs. time-restricted), dietary formulations, and environmental conditions (light intensity and temperature) used in experimental studies as all these factors most likely impact NAD⁺ rhythms.

Redox Status

One of the earliest studies supporting the concept that metabolism can affect clock function investigated the role of cellular redox state (Rutter, et al. 2001). This work showed that DNA binding of the CLOCK-BMAL1 heterodimer could be influenced by the relative ratio of NAD(P)H and NAD(P)⁺. Rutter et al (Rutter et al. 2001) demonstrated that high levels of NAD(P)H increased binding of CLOCK-BMAL1 (or neuronal PAS domain protein 2 (NPAS2)-BMAL1) to DNA E-box motifs, whereas NAD(P)⁺ decreased binding. Notably, high levels of lactate dehydrogenase A (LDHA) mRNA accumulate in cells transfected with expression vectors encoding NPAS2 and BMAL1, demonstrating that LDHA is a transcriptional target of NPAS2-BMAL1. LDH plays a critical role in maintaining cellular redox state as its co-factor for the reaction catalyzing the conversion of lactate to pyruvate (or vice-versa) is NAD⁺ (or NADH). Thus, LDH, a target gene of NPAS2-BMAL1 has the ability to feedback and regulate clock activity by changing the cellular redox potential in vivo. Recent work by Yoshii et al (Yoshii, et al. 2013) confirmed that DNA binding of NPAS2-BMAL1 is enhanced by NAD(P)H, however, NAD(P)⁺ had no effect on DNA binding; these studies also localized the NAD(P)H enhancement binding site to the N-terminal (1–61 residue) region of NPAS2. Together, these studies strongly support the concept that transcriptional activity of the clock is responsive to daily fluctuations in NAD(P)H levels.

Acetylation

The influence of NAD⁺ on the clock is highlighted further by work showing that the NAD⁺-dependent deacetylase SIRT1 impacts clock activity. As discussed in previous sections, intracellular NAD⁺ levels oscillate significantly, due to NAMPT regulation by CLOCK-BMAL1. More recently, evidence suggests that SIRT1 deacetylates BMAL1 and PER2 in an NAD⁺-dependent manner (Asher, et al. 2008; Nakahata, et al. 2008), thereby counteracting the known acetylase activity of CLOCK (which is important for normal clock function) (Doi, et al. 2006). SIRT1 also regulates NAD⁺ levels because it is recruited to the NAMPT promoter (Nakahata et al. 2009a; Ramsey et al. 2009). Therefore, circadian NAD⁺ oscillations influence function of the molecular clock indirectly by controlling rhythms in SIRT1 activity. Evidence exists suggesting that NAD⁺ levels influence the circadian clock in vivo; Sahar et al (Sahar, et al. 2011) reported that CD38 null mice (with chronically elevated NAD⁺ levels) had disrupted circadian behavior (e.g., shortened period length and altered rest-activity rhythms) and altered clock and metabolic gene expression in liver.

ADP-Ribosylation

Another NAD⁺-mediated mechanism for clock regulation has been described, which involves PARP-1 (Asher and Schibler 2011), a protein known to catalyze the transfer of ADP-ribose units from NAD⁺ to various target proteins (Kraus and Hottiger 2013). Asher et al (Asher, et al. 2010) showed that PARP-1 binds to and poly ADP-ribosylates CLOCK, thereby decreasing its binding to DNA. Conversely, loss of PARP-1 enhances CLOCK-BMAL1 binding to DNA, which was found to alter expression of known clock output genes. While PARP-1 activity exhibits diurnal changes, this activity rhythm persists in livers following disruption of the hepatocyte clock, suggesting mediation by extrinsic factors (Asher et al. 2010). Indeed, experiments show that PARP-1 activity is potentially regulated by feeding time, as the peak in poly ADP-ribosylation of PARP-1 can be shifted by restricted feeding protocols (Asher et al. 2010). Therefore, unlike SIRT1, circadian oscillations in NAD⁺ are unlikely to be responsible for PARP-1 activity rhythms, as NAD⁺, NAMPT expression, and PARP-1 activity oscillations are not synchronized, and PARP-1 activity rhythms do not require CLOCK-BMAL1 activity (Asher et al. 2010). Nevertheless, these findings support a mechanism whereby PARP-1 activity links nutritional status to phase shifting of peripheral clocks.

AMP-dependent protein kinase (AMPK)

AMPK is another key metabolic sensor central for transmission of nutrient and energy-dependent signals to the molecular clock (Jordan and Lamia 2013). AMPK functions as an energy sensor, being exquisitely sensitive to the AMP/ATP ratio in cells. When AMP levels are elevated, structural changes in AMPK enhance phosphorylation by liver kinase B1 (LKB1), resulting in activation. AMPK catalyzes the phosphorylation of the clock repressor proteins CRY1 and CRY2, targeting them for proteasomal degradation (Lamia, et al. 2009). AMPK also targets PER proteins for degradation by phosphorylating casein kinase-1ɛ (CK1ε), which subsequently stimulates CK1ε-mediated phosphorylation of PERs (Um, et al. 2007). AMPKα null mice have increased PER2 levels (Um et al. 2007) supporting the hypothesis that AMPK impacts the circadian clock in vivo. Conversely, the AMPK activators AICAR (Um et al. 2007) and metformin (Barnea, et al. 2012) shift the phase of the liver clock. A mechanistic connection also appears to exist between SIRT1 and AMPK; studies show that SIRT1 deacetylates LKB1 influencing the activity, localization, and ability of LKB1 to activate AMPK (Lan, et al. 2008). Collectively, these observations have led to the hypothesis that AMPK may re-set (i.e., entrain) peripheral circadian clocks during specific physiologic and pathophysiologic states.

O-GlcNAcylation

Modification of proteins by O-linked β-N-acetylglucosamine (O-GlcNAc) has emerged as a new regulator linking metabolism to the circadian clock. Reminiscent of phosphorylation, O-GlcNAcylation of proteins is regulated by the balance between O-GlcNAc transferase (OGT; which catalyzes the addition of O-GlcNAc to protein serine and threonine residues), and O-GlcNAcase (OGA; which hydrolyzes O-GlcNAc residues) (Hart, et al. 2011). The substrate for O-GlcNAcylation is the high-energy compound UDP-N-acetylglucosamine; a product of the hexosamine biosynthetic pathway. Studies by Young and colleagues showed coordinated diurnal variations in glucose metabolism, OGT, and OGA leading to peak O-GlcNAcylation of cardiac proteins in the middle of the active (dark) phase (Durgan et al. 2011). These diurnal changes were absent in hearts of cardiomyocyte-specific Clock^?19 mutant mice suggesting these pathways are under direct clock control. Importantly, they demonstrated that BMAL1 is a target for O-GlcNAcylation and that pharmacological inhibition of OGA shifted the phase of the heart clock (Durgan et al. 2011). Related studies in Drosophila and mice also support a role of this PTM in regulation of the circadian clock, as O-GlcNAcylation of CLOCK and PER2 affects their transcriptional activities, and alters the timing of the clock (Kaasik, et al. 2013; Kim, et al. 2012). Furthermore, O-GlcNAcylation prevents BMAL1 and CLOCK degradation by inhibiting their ubiquitination. In addition, liver-specific knockdown of OGT decreased hepatic BMAL1 transcripts late in the active (dark) phase and altered diurnal rhythms in glucose homeostasis (by affecting hepatic gluconeogenic gene expression) (Li, et al. 2013). Together, these studies demonstrate that O-GlcNAc is novel metabolic signal connecting metabolism and the circadian clock.

Heme and Reactive Species

The iron-containing protein heme influences clock function by binding to distinct clock components, including CRY1–2, NPAS2, and REV-ERBα (Burris 2008). Heme binding to REV-ERB nuclear receptors causes the recruitment of the nuclear receptor co-repressor (NCoR)- histone deacetylase (HDAC) co-repressor complex to REV-ERB homodimers leading to repression of target genes (e.g., BMAL1) (Raghuram, et al. 2007; Yin and Lazar 2005). The rate limiting enzyme in heme biosynthesis, δ-aminolevulinate synthase (ALAS1), exhibits a circadian rhythm at the gene level (Rogers, et al. 2008) due to activation by NPAS2 and PGC1α, resulting in circadian clock mediated oscillations in heme levels (Handschin, et al. 2005; Kaasik and Lee 2004). Importantly, ligation of heme to REV-ERBα in turn inhibits its own biosynthesis through REV-ERBα-mediated inhibition of PGC-1α (Wu, et al. 2009). This feedback loop functions to tightly maintain heme levels within a physiologic range in the cell. Thus, heme metabolism appears to constitute an additional metabolic feedback loop of the molecular clock.

An important characteristic of heme is its ability to bind gaseous signaling molecules including nitric oxide (NO), carbon monoxide (CO), and hydrogen sulfide (H₂S). The ability of gaseous signaling molecules to influence clock function was first recognized when binding of CO to the heme in NPAS2 decreased formation of NPAS2-BMAL1 heterodimers in vitro (Dioum, et al. 2002). Transcriptional regulation by REV-ERBs may be similarly influenced by interaction of heme with these gaseous molecules. Pardee et al (Pardee, et al. 2009) reported that the NO donor (DETA NONOate) blocked REV-ERB mediated repression of gene transcription in HEK 293 cells. Work by Gupta and Ragsdale have expanded on this novel concept by showing that the redox state of a thiol-disulfide redox switch controls heme affinity to the ligand binding domain in REV-ERBβ (Gupta and Ragsdale 2011). This study demonstrated that formation of a disulfide bond between Cys-384 and Cys-374 in the REV-ERBβ ligand binding domain lowers the affinity for heme, whereas the reduced dithiol between these two Cys residues binds heme more tightly. As thiol-disulfide switches are sensitive to PTM by reactive oxygen and nitrogen species (ROS and RNS), conditions involving oxidative stress may cause perturbation of REV-ERB controlled pathways including the clock. With regards to H₂S, much less is known about the impact this signaling molecule may have on the molecular clock. Incubation of cultured mouse hepatocytes with sodium hydrosulfide (NaHS) was shown to maintain oscillations of distinct clock genes (Shang, et al. 2012). Furthermore, inhaled H₂S was found to increase cardiac PER1 gene expression relative to rats exposed to room air (unpublished results). Together, the results from these studies strongly suggest that interaction of heme with multiple reactive gaseous species may be important for maintenance of clock function, whereas aberrant production of reactive species could potentially dysregulate the clock. The importance of this level of control is strengthened further by recent reports that peroxiredoxin, the main hydrogen peroxide detoxification enzyme, has a 24 hour cycle in the oxidation state of its catalytic site (Edgar, et al. 2012). Indeed, future studies should be aimed at understanding circadian regulation of the enzymatic pathways responsible for the production and removal of the gaseous signaling molecules and other reactive species (e.g., ROS) as mounting evidence support their role in clock function.

Shift Work

Night-shift workers often exhibit dyslipidemia (e.g., high triglycerides and/or low HDL cholesterol), increased postprandial serum glucose and insulin (Lund, et al. 2001), and increased circulating levels of several biomarkers of inflammation (Sookoian, et al. 2007), associated with increased incidence of obesity and diabetes (Burgueno, et al. 2010; Chen, et al. 2010; Copertaro, et al. 2008; De Bacquer, et al. 2009; Karlsson, et al. 2001; Kroenke, et al. 2007; Lin, et al. 2009; Morikawa, et al. 2005; Nagaya, et al. 2002). Controlled laboratory-based studies clearly showed that circadian misalignment disrupts metabolism in humans. For example, Scheer et al (Scheer et al. 2009) induced circadian misalignment in healthy adults by using an 11-day forced dyssynchrony protocol consisting of repeated ‘28 hr days’; during this protocol, all subjects ate four isocaloric meals each ‘28 hr day’ and slept at all phases of the circadian cycle. The results showed that circadian misalignment (defined as eating and sleeping 12 hour out of phase from their habitual times) increased circulating glucose and insulin, decreased leptin, and increased blood pressure. Indeed, some subjects in this study exhibited pre-diabetic symptoms in response to circadian misalignment. More recently, Van Cauter and colleagues reported augmentation of insulin resistance in sleep deprived subjects following circadian misalignment (Leproult, et al. 2014). Together, these results support the concept that circadian misalignment in shift workers may contribute towards cardiometabolic disease development.

The influence of sleep disturbances (e.g., too little sleep) and/or psychosocial stressors should not be overlooked as additional risk factors for metabolic disease in shift workers (Bannai and Tamakoshi 2014; Clougherty, et al. 2010). For example, a combination of sleep deprivation and circadian disruption (‘28 hr days’) caused decreased metabolic rate and increased postprandial glucose in healthy adults (Buxton, et al. 2012). Sleep deprivation and sleep restriction alone have been shown to alter endocrine regulators of glucose and energy metabolism, which may lead to insulin resistance, obesity, and other pathologies (Lucassen, et al. 2012; Zizi, et al. 2010). Studies show that acute sleep restriction (3 consecutive nights of shortened sleep – 4 hr of sleep per night) reduced insulin sensitivity in adolescent boys (Klingenberg, et al. 2013). Schmid et al (Schmid, et al. 2007) reported a significant decrease in plasma glucagon levels following one night of total sleep deprivation in healthy young men aged 20–40 year. Interestingly, plasma glucagon levels were significantly decreased in the morning following only one night of mild sleep restriction (one night of 4.5 hour of sleep) as compared to levels measured in healthy men allowed to sleep for 7 hour (Schmid, et al. 2009). In this study, plasma glucagon levels also remained depressed throughout a 240 minute step-wise hypoglycemic clamp in the sleep restricted group. Sleep deprivation (24 hour of continual wakefulness) significantly decreased energy expenditure (resting and postprandial), increased morning plasma glucose (postprandial), and elevated morning levels plasma ghrelin levels, and increased nighttime and daytime levels of cortisol as compared to the control group that was allowed 8 hour of sleep during a normal 24 hour sleep-wake cycle (Benedict, et al. 2011). While the mechanisms responsible for these effects remain unknown, they provide compelling evidence to support a linkage among disturbed sleep, dysregulated glucose and energy metabolism, and a predisposition to adverse metabolic consequences.

Experimental animal models have also been utilized to demonstrate that circadian misalignment causes metabolic disturbances. Using a ‘night-work’ model in rats, Salgado-Delgado et al (Salgado-Delgado, et al. 2010) showed that forced activity (and/or food consumption) during the normal rest phase increased body weight, adiposity, and glucose intolerance compared to rats with forced activity in the normal active phase. Further, this experimental protocol disrupted the normal daily rhythms in both clock and metabolic genes, and caused hepatic steatosis (Salgado-Delgado, et al. 2013). Similarly, dim-light-at-night (dLAN) has been shown in increase weight gain and disrupt circadian rhythms in both the SCN and peripheral tissues (Fonken, et al. 2013a; Fonken, et al. 2010), as well as increase peripheral inflammation when combined with a high fat diet (Fonken, et al. 2013b). Interestingly, re-exposure to a dark night can reverse metabolic disturbances following exposure to dLAN (Fonken, et al. 2013c). These data support the hypothesis that activity and/or light in the normal rest (or sleep) phase causes misalignment among different organ systems and intracellular metabolic pathways. Accordingly, circadian misalignment or dyssynchrony may partially explain why night- and shift-work is strongly associated with obesity and related metabolic diseases. An additional cause of circadian misalignment during shift work is potentially the timing of food intake (as discussed below).

Timing of Feeding

Food consumption is a potent zeitgeber for peripheral circadian clocks, as first demonstrated by Damiola et al; restricting food access to the sleep phase shifted circadian clocks in the liver, kidney, and heart, with no effect in the SCN, thereby result in circadian dyssynchrony between central and peripheral clocks (Damiola et al. 2000). More recently, studies report that this experimental protocol differentially influences clocks within metabolically active tissues (e.g., liver, skeletal muscle, and adipose), suggesting that meal feeding also causes dyssynchrony between peripheral clocks (Bray et al. 2013). This is associated with perturbations in metabolic homeostasis, resulting in increased adiposity, decreased glucose tolerance, and dyslipidemia, a metabolic profile often observed in night-eating syndrome subjects (Arble, et al. 2009; Bray et al. 2013; Stunkard and Allison 2003). Conversely, restricting food intake to the normal activity phase in rodents prevents metabolic disturbances and weight gain associated with high fat diets (Hatori, et al. 2012; Tsai, et al. 2013). Evidence also suggests that the timing of macronutrient ingestion within the active/awake period markedly impacts metabolic outcomes. More specifically, consumption of a calorically dense high fat meal at the end of the active/awake period increases adiposity, decreases glucose tolerance, and increases circulating insulin and leptin levels in mice (relative to consumption of the exact same meal during the beginning of the active period) (Bray, et al. 2010). This experimental paradigm phase shifts peripheral circadian clocks (unpublished observations). Interestingly, when human subjects were forced to consume the majority of their calories immediately prior to overnight sleep, metabolic perturbations were observed (e.g., dissociation between circulating insulin and glucose levels) (Qin, et al. 2003). Collectively, these observations suggest that consumption of nutrients at an inappropriate time of the day can result in circadian dyssynchrony, associated with metabolic imbalances.

Alcohol Consumption

In addition to high fat diets, recent studies reported that other life-style factors like chronic alcohol consumption can alter circadian rhythms in peripheral tissues like gut (Forsyth, et al. 2013; Summa, et al. 2013; Swanson et al. 2011) and liver (Filiano et al. 2013; Zhou, et al. 2014); thus, contributing to circadian misalignment and tissue injury. Work from our laboratories revealed that diurnal oscillations in clock genes and clock-controlled genes were altered in liver of ethanol-fed mice exhibiting steatosis (Filiano et al. 2013). Importantly, ethanol feeding induced a phase advance in clock gene expression in liver, but not in SCN, indicating dyssynchrony between the master clock in the SCN and the peripheral clock in the liver. Ethanol consumption also altered the diurnal rhythms in several lipid, carbohydrate, and energy metabolism genes (Filiano et al. 2013), which were confirmed by (Zhou et al. 2014). These findings are important because they revealed that chronic alcohol consumption perturbs the circadian clock in the liver and associated downstream metabolic targets. Notably, these results support the hypothesis that ethanol, like high fat, induces circadian misalignment between molecular oscillators. As a consequence, the peripheral liver clock is no longer in sync with its environment during alcoholic and nonalcoholic fatty liver disease.

Obesity/Diabetes

It is clear that circadian dyssynchrony negatively impacts cardiometabolic parameters. What has also become apparent is that cardiometabolic diseases result in circadian dyssynchrony. For example, Young et al (Young, et al. 2002) reported that circadian clock gene expression was phase shifted in the rat heart following streptozotocin (STZ)-induced diabetes, an effect that has been confirmed for various peripheral clocks (Oishi, et al. 2004). Several classic paramaters of clock gene expression (e.g., acrophase, amplitude, and mesor) were found to be altered in liver of out-bred male rats treated with STZ or in liver of male spontaneous type 1 diabetes LEW.1AR1-iddm rats (Hofmann, et al. 2013). Specifically, the acrophase of BMAL1 and other clock component genes were phase advanced in liver of STZ-treated and iddm rats, with insulin treatement normalizing some of the dysregulated clock rhythms. Furthermore, peripheral clocks also appear to be phase shifted in rodent models of type 2 diabetes (e.g., db/db mice) (Kudo, et al. 2004). Diet-induced obesity, from unrestricted access to a high fat diet, altered rhythms in circadian behaviors (activity and feeding), neuropeptides in the hypothalamus, and clock and metabolism gene expression in liver and adipose tissue of mice (Kohsaka et al. 2007). Other studies have reported similar findings in response to diet-induced obesity in highly metabolic tissues (Bravo, et al. 2014; Lizier, et al. 2013; Pendergast, et al. 2013; Prasai, et al. 2013; Shi, et al. 2013). Recent work by Sassone-Corsi and colleagues also shows that a high fat diet disrupts normal circadian oscillations of the molecular clock and clock-controlled metabolites and transcripts in the liver (Eckel-Mahan, et al. 2013); they propose that these high fat diet-mediated alterations in metabolism are linked to impaired CLOCK-BMAL1 chromatin recruitment and induction of peroxisome proliferator-activated receptor gamma (PPARγ)-mediated oscillations in noncyclic genes (Eckel-Mahan et al. 2013). Taken together, these studies support the hypothesis that circadian misalignment consequent to environmental, life-style, and dietary factors most likely contributes to the development of cardiometabolic diseases.