The Circadian Clock Coordinates Ribosome Biogenesis

We investigated whether the circadian clock might coordinate translation in mouse liver. Indeed, quantitative reverse transcription (RT)-PCR analyses confirmed that mRNAs of most of the factors involved in translation initiation are rhythmically expressed with a period of 24 h (Figure 1A; statistical analyses are given in Table S1). Interestingly, while we did not observe any significant variations in protein abundance, rhythmic phosphorylations were strongly manifested during two consecutive days, emphasizing the robustness of these rhythms (Figure 1B; quantification and statistical analyses of the data are given on Figure S1 and Table S2). EIF4E is mostly phosphorylated during the day, with a peak at the end of the light period (ZT6-12), whereas EIF4G, EIF4B, 4E-BP1, and ribosomal protein (RP) S6 (RPS6) are mainly phosphorylated during the night, which is, in the case of nocturnal animals like rodents, the period when the animals are active and consume food.

Phosphorylation of these factors is well characterized and involves different signaling pathways [8] whose reported activity perfectly correlates with the observed phosphorylation rhythm. EIF4E is phosphorylated by the extracellular signal-regulated protein kinase (ERK)/mitogen-activated protein kinase (MAPK)-interacting kinase (MNK) pathway [9], which is most active during the day, at the time when EIF4E reaches its maximum phosphorylation (Figure 2A; quantification and statistical analyses of the data are given on Figure S2 and Table S2). On the other hand, EIF4G, EIF4B, 4E-BP1, and RPS6 are mainly phosphorylated by the target of rapamycin (TOR) complex 1 (TORC1) [10], which is activated during the night, at the time when the phosphorylation of these proteins reaches its maximum level. TORC1, in turn, is negatively regulated by the tuberous sclerosis protein complex (TSC), whose activity is under the control of the phosphoinositide 3-kinase (PI3K)/AKT, ERK, and the energy sensing 5′ adenosine monophosphate-activated protein kinase (AMPK) pathways [10],[11]. As reported [12], AMPK is active during the day and mediates the activation of TSC2, contributing to the repression of TORC1 in the period of energy and nutrient restriction. Conversely, during the night, TORC1 is activated probably through TSC2 inhibition by PI3K via TORC2 [13].

Interestingly, we found that mTor, its partner Raptor, as well as its regulating kinase Map3k4, are also rhythmically expressed, thus potentially further contributing to the rhythmic activation of TORC1 (Figure S3; Table S1). ERK is activated during the day in synchrony with the rhythmic expression of Mnk2 (Figure S3), contributing to EIF4E phosphorylation during this period. However, its downstream target RPS6 Kinase (RSK) seems to contribute only marginally to the phosphorylation of RPS6 in mouse liver (Figures 1B and 2A). The rhythmic phosphorylation of 4E-BP1 resulted in its release from the mRNA cap-mimicking molecule 7-methyl-GTP from ZT14 to ZT22 (Figure 2B; Table S2), allowing the rhythmic assembly of the EIF4F and potentially mRNA translation.

The rhythmic expression of mRNA encoding translation initiation factors, TORC1 complex component, and a kinase activating these factors is independent of light as it is maintained under constant darkness, even if the phase seems to be advanced (Figure S4A). Interestingly, activation of the TORC1 pathway is also maintained under constant darkness but with an advanced phase (Figure S5A). Since nutrient availability is a potent activator of the TORC1 pathway [13], we asked whether these parameters are also rhythmic under conditions of starvation. We found that expression of mRNA encoding translation initiation factors, TORC1 complex component, and a kinase activating these factors is still rhythmic under starvation (Figure S4B), even when this starvation occurs under constant darkness (Figure S4C). This result unambiguously demonstrates the role of the circadian clock in the expression of these genes. In addition, phosphorylations of RPS6 and 4E-BP1 are still rhythmic under starvation, whether or not the mice are under a light-dark regimen or in constant darkness (Figure S5B and S5C), confirming previously published observations [14]. Interestingly, TORC1 activation is in opposite phase with the clock-dependent rhythmic activation of autophagy in mouse liver [15], a process inhibited by TORC1 but able to generate amino acids that can in turn activate TORC1 [16]. This might suggest that the circadian clock can regulate the two processes in a coordinated fashion. Importantly, rhythmic activation of TORC1 is not restricted to the liver as the same phosphorylation rhythm is found in kidney and heart, albeit with reduced amplitude (Figure S6). Meanwhile, TORC1 activation is constant in brain, lung, and small intestine, suggesting that the rhythmic nutrient availability due to the circadian clock-regulated feeding behavior is not sufficient by itself to explain the rhythmic activation of TORC1.

Diurnal binding of 4E-BP to EIF4E suggested that translation might be rhythmic in the liver. To test this hypothesis and to identify potential rhythmically translated genes, we purified polysomal RNAs, a RNA sub-fraction composed mainly of actively translated mRNA, every 2 h during a period of 48 h. We found that relative amount of this polysomal fraction follows a diurnal cycle, showing that a rhythmic translation does occur in mouse liver (Figure S7). This result confirms original observations based on electron microscopy and biochemical studies [17],[18]. We therefore decided to characterize these rhythmically translated mRNAs through comparative microarray analysis of polysomal and total RNAs. While the obtained profiles in polysomal and total RNAs fractions are highly similar for most mRNAs (examples of rhythmic mRNAs are given on Figure S8), 249 probes showed a non-uniform ratio in diurnal polysomal over total mRNAs (Figure 3A). This means that approximately 2% of the expressed genes are translated with a rhythm that is not explained by rhythmic mRNA abundance as in most cases, the total mRNA levels were constant while the polysomes-bound mRNA levels fluctuated during the 24-h cycle (Figures 3B and S9). Among translationally regulated genes, 70% were found in the polysomal fraction during the same time interval, starting at ZT8 before the onset of the feeding period and finishing at the end of the dark period (Tables S3 and S4). Most of these genes belonged to the 5′-terminal oligopyrimidine tract (5′-TOP) family, known to be regulated by TORC1 [19], but also by the level and phosphorylation state of EIF4E [20],[21]. 5′-TOP genes are themselves involved in translation via ribosome biogenesis and translation elongation (Table S4).

After confirmations of these results by quantitative RT-PCR (Figure S10), we wished to validate the periodicity in the amount of mRNAs purified in the different fractions obtain during polysomes purification over a 24-h period. Whereas a constitutively translated mRNA such as Gapdh is found all the time in the polysomal fraction (with a small decrease in the middle of the light period when overall translation decreases), mRNAs coding for RPs are associated with the polysomal fraction only starting towards the end of the light period (ZT8) and during the dark period (Figure 3C). This result demonstrates a dynamic translation initiation of 5′-TOP mRNA starting before the onset of the feeding period, with a maximum at the beginning of the dark period.

Next, we wanted to confirm that this rhythmic translation had an impact on the protein levels. With respect to RPs, while the half-life of mature ribosomes is approximately 5 d in rodent liver [22], newly synthesized RPs have a half-life of only a few hours, as most of them are rapidly degraded after translation during the ribosome assembly process in the nucleolus [23]. We thus expected a rhythmic expression of this subpopulation of newly synthesized RPs in the soluble cytosolic fraction depleted of ribosomes after sedimentation. Indeed, under these conditions, RPs show a rhythmic abundance with highest expression during the night (Figure 3D; quantification and statistical analyses of the data are given on Figure S11 and Table S2). In some cases, we noticed a shallow decrease at ZT16-18, potentially reflecting transport of RPs into the nucleolus for ribosome assembly.

In addition to translational regulation, we also observed a diurnal expression of RP mRNAs, albeit with a small average peak to trough amplitude of approximately 1.2. Taking into account their relatively long half-life (11 h) [24], we hypothesized that this minor fluctuation might reflect more pronounced rhythmic amplitudes in transcription as amplitude decreases with half-life [25]. In addition, it has recently been shown that the transcription of several RP mRNAs is directly controlled by the molecular oscillator in Drosophila head [26]. Indeed, pre-mRNA accumulation of several RP exhibited a rhythmic transcription, with an average amplitude of 3.5-fold with a maximum at ZT8, just before the activation of their translation (Figure 4A; statistical analyses are given in Table S1). In addition, we found that the synthesis of the ribosome constituent precursor 45S rRNA is also rhythmic and synchronized with RP mRNAs transcription, indicating that all elements involved in ribosome biogenesis are transcribed in concert, then translated or matured. In yeast [27] and Drosophila[28], transcription of RP mRNAs appears to be coordinated with rRNA transcription, which is a rate limiting step in ribosome biogenesis. On the other hand, in mammals, rRNA transcription is highly regulated by the upstream binding factor (UBF), which establishes and maintains an active chromatin state [29]. Remarkably, we found that UBF1 is rhythmically expressed in mouse liver at both mRNA and protein levels (Figure 4B; quantification and statistical analyses of the data are given in Figure S12A and Tables S1 and S2), in phase with RP mRNAs and rRNAs transcription. In addition, rhythmic transcription of Ubf1 and Rpl23 genes is also independent of light and food (Figure S4).

To test whether Ubf1 transcription is regulated by the circadian clock, we characterized its expression in arrhythmic Cry1/Cry2 knockout (KO) [30] and Bmal1 KO [31] mice, which are devoid of a functional circadian clock. Indeed, these mice do exhibit an arrhythmic pattern of activity under constant darkness, which is in general correlated with an arrhythmic feeding behaviour. As TORC1, as well as other signaling pathways, are in part regulated by feeding through nutrient availability, we expect a temporally discontinuous and erratic activation of these pathways in the KO mice under unrestricted feeding. To verify this hypothesis, we measured activation of the TORC1, AKT, and ERK pathways in Cry1/Cry2 and Bmal1 KO kept in constant darkness. As shown in Figure S13A, the rhythmic activation of these signaling pathways is indeed lost under this condition, confirming their arrhythmic activation. To highlight the role of the feeding regimen on this activation, we kept Cry1/Cry2 KO mice in constant darkness and sacrificed them at CT12. We found a strong inter-individual variability in the activation of the TORC1, AKT, and ERK pathways, reflecting the arrhythmic feeding rhythm of these animals (Figure S13B). To circumvent this caveat and study the rhythmic translation in mice devoid of a functional molecular oscillator, we decided to place Cry1/Cry2 and Bmal1 KO under a light-dark regimen to keep a normal diurnal feeding behaviour due to masking. In addition, mice had access to food only during the dark phase to eliminate the effect of a potential disturbed feeding behaviour. Under these conditions, KO mice had a rhythmic feeding behaviour and thus potential differences in protein levels or pathway activity cannot be attributed to the arrhythmic feeding behaviour of these animals. We indeed found that UBF1 rhythmic expression is dependent on a functional circadian clock as it is impaired in both animal models (Figure 4C and 4D; quantification and statistical analyses of the data are given in Figure S12B and Tables S5, S6, S7, S8). However, if UBF1 expression is persistently low in Cry1/Cry2 KO mice, this expression is constantly high in Bmal1 KO mice, suggesting the control of Ubf1 by a circadian clock-regulated transcription repressor. In addition, we observed that these animals lose also the synchrony and coordination of 45S rRNA and RP pre-mRNAs transcription (Figures 5, S14, and S15; statistical analyses of the data are given in Table S5 and S6). Indeed, decreased UBF1 expression in Cry1/Cry2 KO mice is correlated with lower 45S rRNA transcription, but higher and delayed RP pre-mRNAs transcription. Interestingly, Bmal1 KO mice present a complete arrhythmic transcription of RP pre-mRNAs, highlighting the crucial role of the circadian clock in the coordination of rRNA and RP mRNAs transcription.

Rhythmic expression of genes coding for components of the translation initiation complex is strongly dampened or phase-shifted in both KO models, in addition to an altered level of expression (Figures 5, S14, and S15; statistical analyses of the data are given in Tables S5 and S6). However, we did not observe in general any significant variations in protein abundance, excepting a slight increase in EIF4E expression in Cry1/Cry2 KO mice, reflecting increased mRNA expression (Figure 6A and 6C; quantification and statistical analyses of the data are given in Figures S16, S17; Tables S7 and S8). The variations in EIF4G levels reflect more the changes in its phosphorylation state, which regulates its stability [32]. While most of the signaling pathways are still rhythmic in Cry1/Cry2 KO mice, except for the ERK pathway and the downstream phosphorylation of EIF4E, which loses its rhythmic activation, the phase of the activation of the TORC1 and AKT pathways are advanced in comparison to wild-type (WT) mice (Figures 6A and S16; quantification and statistical analyses of the data are given in Table S7). As a consequence, the rhythmic expression of RPs is altered in Cry1/Cry2 KO mice (Figure 6B; quantification and statistical analyses of the data are given in Table S7), with an increased level of expression, likely because of the increased RP pre-mRNAs and EIF4E levels [20], and a delayed phase of expression. Most of the rhythmic activation of the three pathways is also strongly altered in Bmal1 KO mice (Figures 6C and S17; quantification and statistical analyses of the data are given in Table S8). As shown in Figure 6D, the phase of RPs rhythmic expression is severely advanced with a maximum of expression in the middle of the day instead of the night (Figure 6D; quantification and statistical analyses of the data are given in Table S8).

The results presented here show that the molecular circadian clock controls ribosome biogenesis through the coordination of transcriptional, translational, and post-translational regulations. Moreover, the data strongly suggest that a functional molecular oscillator is required for a timely coordinated transcription of translation initiation factors, RP mRNAs, and rRNAs. The clock modulates the rhythmic activation of signaling pathways controlling translation through the TORC1 pathway, translation of RPs, and ribosome biogenesis (Figure 7). Interestingly, it has been reported that the size of the nucleolus, the site of rRNA transcription and ribosome assembly, follows a diurnal pattern with a maximum in the middle of the dark period [33], which thus occurs in synchrony with the observed accumulation of RPs in the liver. The observed rhythmic ribosome biogenesis is substantiated by the previous observation showing that both size and organization of the nucleolus are directly related to ribosome production [34].

Remarkably, a coordinated rhythmic regulation of transcriptional and translational events for the biogenesis of ribosomes has also been suggested for the filamentous fungus Neurospora crassa[35] and for plants [36],[37]. Since ribosome biogenesis is one of the major energy consuming process in cells [38], its tight control is primordial to reduce interferences with other biological processes. In the case of mouse liver, we estimate that the decrease of translation during the light period is equivalent to 20% of the total translation (Figure S7), in agreement with previously published results [17]. Although moderate, this decrease affects translation of housekeeping genes like Gapdh (Figure 3C) and probably the translation of other genes. It means that the increase in ribosome biogenesis during the night could potentially influence the translation of many other mRNAs, however with a magnitude sufficiently low to not allow its detection by our method.

Nevertheless, it is clear that this energy-consuming process has to be confined to a time when energy and nutrients are available in sufficient amount, which, in the case of rodents, is during the night period when the animals are active and consume food. Hence, all the elements required for translation have to be ready to start ribosome biogenesis during that time. This is achieved by increasing levels of rRNAs and RP pre-mRNAs just before the onset of the night, synchronized with the phosphorylation of EIF4E that increases 5′-TOP mRNAs translation [21]. Activation of the TORC1 pathway during this period promotes RPs synthesis, rRNAs maturation, and ribosome assembly. In addition activation of the ERK pathway correlates also with ribosome biogenesis [39], strengthening the rhythmic nature of this process. Accordingly, orchestration of ribosome biogenesis by the circadian clock represents a nice example of anticipation of an obligatory gated process through a complex organization of transcriptional, translational, and post-translational events.

As described in the introduction, the mammalian molecular circadian oscillator consists in interlocked feedback loops of transcription factors that generate a complex network of rhythmically expressed genes [3]. Within the core molecular clock, increasing evidence shows that post-translational modifications play a crucial role in the generation of circadian rhythms [40]. However, the circadian clock is also able to coordinate rhythmic post-translational activation of signaling pathways not directly involved in the molecular oscillator but rather in the sensing of the environment. The first described example consisted in the rhythmic activation of ERK in the suprachiasmatic nucleus (SCN) of the hypothalamus where the master circadian pacemaker is localized: if light stimulates ERK phosphorylation in the SCN in a time-dependent fashion, circadian ERK phosphorylation continues also in constant darkness, suggesting a crucial role of the circadian clock in this process [41]. Interestingly, the same observations have been made for the TORC1 pathway in the SCN [42],[43], and for the PI3K/AKT pathway in the retina [44]. Considering the fact that these two pathways have been recently identified as a potent regulators of circadian activity in Drosophila[45], we expect that the role of the circadian clock-coordinated signaling pathways on circadian physiology will probably be emphasized in other organisms in the near future.

With respect to rhythmic activation of signaling pathways in the liver, there are only few examples of such regulations. One example is the rhythmic activation of the PI3K/AKT pathway that is associated with food metabolism and rhythmic feeding behavior [46]. Recently, we also described a circadian clock-dependent rhythmic activation of the unfolded protein response regulating liver lipid metabolism [47]. In addition, it has been shown that the circadian clock is also able to regulate autophagy in mouse liver [15]. In this context, our discovery of the rhythmic ribosome biogenesis through coordination of the rhythmic activation of signaling pathways constitutes an important new element in this area of research.

It has long been known that caloric restriction or intermittent fasting increases lifespan in a wide variety of models [48]. Increased lifespan has also been linked to the reduced activation of the TORC1 pathway, which, in turn, provokes a reduced mRNA translation [49],[50]. The role of the TORC1 pathway in this translation-dependent extension of lifespan has been genetically confirmed in Caenorhabditis elegans[51] and Drosophila[52],[53]. A similar scenario is also considered in mice since treatment with the TOR inhibitor rapamycin [54] or deletion of the TORC1 downstream protein kinase S6K1 [55] lead to increased lifespan. In addition, downregulation of various components of the EIF4F complex extends lifespan in C. elegans[56]–[59], whereas inhibition of RPs genes expression extends lifespan in both Saccharomyces cerevisiae[60] and C. elegans[56]. Hence, keeping ribosome biogenesis, and translation in general, to their minimum levels plays a major role in the regulation of longevity [61]. Interestingly, all the genetically modified animal models presenting a disrupted circadian clock [62]–[64] or mice subjected to chronic jet lag [65] are subjected to premature aging and reduced lifespan. The deregulation of many other circadian-clock regulated processes can reduce life expectancy, like reduced xenobiotic detoxification [66]. We thus believe that the potential role of disorganized ribosome biogenesis on life expectancy, observed in animals devoid of a circadian clock, will be an exciting subject for further studies.

All animal studies were conducted in accordance with our regional committee for ethics in animal experimentation and the regulations of the veterinary office of the Canton of Vaud. C57Bl/6J mice were purchased from Janvier (Le Genest) or Charles River Laboratory (L'Arbresle). Bmal1 floxed mice have been previously described [67]. These mice were crossed with mice expressing the CRE recombinase under the control of the CMV promoter [68] to obtain Bmal1 KO mice. Cry1/Cry2 double KO mice [30] in the C57Bl/6J genetic background have been previously described [69]. In all experiments, male mice between 10 and 12 wk of age are used. Unless noted otherwise, mice were maintained under standard animal housing conditions, with free access to food and water and in 12-h light/12-h dark cycles. However, for all experiments, animals were fed only at night during 4 d before the experiment to reduce effects of feeding rhythm. For experiments in constant darkness, mice were shifted into complete darkness after the last dark period and then sacrificed every 2 or 4 h during the next 48 h. For starvation experiments, mice were deprived from food during one complete night and then during the following 24 h, mice were sacrificed every 2 or 4 h.

Livers were homogenized in lysis buffer containing 20 mM HEPES (pH 7.6), 250 mM NaCl, 10 mM MgCl₂, 10 mM DTT, 20 µg/ml cycloheximid, 10 U/µl RNase inhibitor, and a protease inhibitor cocktail containing 0.5 mM PMSF, 10 µg/ml Aprotinin, 0.7 µg/ml Pepstatin A, and 0.7 µg/ml Leupeptin. The homogenates were centrifuged 10 min at 9,500 g and 1 mg/ml heparin, 0.5% Na deoxycholate, and 0.5% Triton ×100 were added to the supernatant. 50 mg of lysate were deposited on a 36 ml 7% to 47% sucrose gradient in a buffer containing 20 mM HEPES (pH 7.6), 100 mM KCl, 5 mM MgCl₂, and 1 mM DTT. After 4 h 30 min of centrifugation at 130,000 g and 4°C, the gradient was divided in fractions of approximately 1 ml with a peristaltic pump. Optic density of the fractions at 260 nm was measured to establish the polysomal profile in the gradient. Fractions were finally pooled in ten fractions. An example of polysome profile is given on Figure S18. RNAs were then extracted according to the protocol described by Clancy et al. [70] that we slightly modified. Briefly, fractions were precipitated by the addition of three volumes of ethanol and kept overnight at −80°C. After 30 min of centrifugation at 5,200 g, RNAs were extracted from the non-soluble fraction by classical protocol [71].

Liver RNAs were extracted and analysed by real-time quantitative RT-PCR, mostly as previously described [25]. Briefly, 0.5 µg of liver RNA was reverse transcribed using random hexamers and SuperScript II reverse transcriptase (Life Technologies). The cDNAs equivalent to 20 ng of RNA were PCR amplified in triplicate in an ABI PRISM 7700 Sequence Detection System (Applied Biosystem) using the TaqMan or the SYBR Green technologies. References and sequences of the probes are given in Tables S9 and S10, respectively. Gapdh mRNA (total RNA) or 28S rRNA (polysomal RNA) were used as controls.

Liver polysomal and total RNAs were extracted independently from two mice sacrificed every 2 h during 48 h. For polysomal RNAs, we pooled fractions 1 and 2 from the ten fractions obtained during the extraction and containing heavy polysomes. 3 µg of polysomal and total RNAs from each animal from each time point were pooled. These 6 µg of polysomal and total RNAs were used for the synthesis of biotinylated cRNAs according to Affymetrix protocol, and hybridized to mouse Affymetrix Mouse Genome 430 2.0 arrays. The chips were washed and scanned, and the fluorescence signal analysed with Affymetrix software. Data are deposited on the Gene Expression Omnibus database under the reference GSE33726↗ (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?token=rpwvtoqogkamwrm&acc=GSE33726↗).

The raw data of all 48 arrays were normalized together using the robust multiarray average (RMA) method [72]. For the analysis, we filtered out all probesets corresponding to introns using the Ensembl annotation and then only kept genes with a sufficient expression level (we kept genes whose probe signal in the total fraction was above 5 in log2 scale). For the identification of circadian probesets, the 24-h Fourier component (F24) and the phase were computed using established methods [73]. The associated p-value (p) was calculated using the Fisher test (p = (1−s)¹⁰) [73]. For the identification of rhythmically translated genes, the difference between polysomal and total RNAs was subjected to Fourier analysis and we selected probesets giving a p-value inferior to 0.001. In addition, we requested that the peak to trough amplitude in the polysomal signal be above 1.2-fold.

Nuclear and cytoplasmic proteins were extracted mostly as described [25]. Briefly, liver were homogenized in sucrose homogenization buffer containing 2.2 M sucrose, 15 mM KCl, 2 mM EDTA, 10 mM HEPES (pH 7.6), 0.15 mM spermin, 0.5 mM spermidin, 1 mM DTT, and the same protease inhibitor cocktail as for polysomes extraction. Lysates were deposited on a sucrose cushion containing 2.05 M sucrose, 10% glycerol, 15 mM KCl, 2 mM EDTA, 10 mM HEPES (pH 7.6), 0.15 mM spermin, 0.5 mM spermidin, 1 mM DTT, and a protease inhibitor cocktail. Tubes were centrifuged during 45 min at 105,000 g at 4°C. After ultra-centrifugation, supernatants containing soluble cytoplasmic proteins were harvested, homogenised, and centrifuged for 2 h at 200,000 g to remove ribosomes. These supernatants constitute cytoplasmic extracts. The nucleus pellets were suspended in a nucleus lysis buffer composed of 10 mM HEPES (pH 7.6), 100 mM KCl, 0.1 mM EDTA, 10% Glycerol, 0.15 mM spermine, 0.5 mM spermidine, 0.1 mM NaF, 0.1 mM sodium orthovanadate, 0.1 mM ZnSO4, 1 mM DTT, and the previously described protease inhibitor cocktail. Nuclear extracts were obtained by the addition of an equal volume of NUN buffer composed of 2 M urea, 2% nonidet P-40, 600 mM NaCl, 50 mM HEPES (pH 7.6), 1 mM DTT, and a cocktail of protease inhibitor, and incubation 20 min on ice. After centrifugation during 10 min at 21,000 g, the supernatants were harvested and constitute nuclear extracts.

25 µg of nuclear or 12.5 µg cytoplasmic extracts were used for western blotting. After migration, proteins were transferred to PVDF membranes and Western blotting was realized according to standard procedures. References for the antibodies are given in. Table S11

Organs were homogenized in lysis buffer containing 20 mM HEPES (pH 7.6), 100 mM KCl, 0.1 mM EDTA, 1 mM NaF, 1 mM sodium orthovanadate, 1% Triton X-100, 0.5% Nonidet P-40, 0.15 mM spermin, 0.5 mM spermidin, 1 mM DTT, and a protease inhibitor cocktail. After incubation 30 min on ice, extracts were centrifuged 10 min at 21,000 g and the supernatants were harvested to obtain total extracts.

65 µg of extract was used for Western blotting. After migration, proteins were transferred to PVDF membranes and Western blotting was realized according to standard procedures. References for the antibodies are given in. Table S11

7-methyl GTP sepharose 4B beads (GE Healthcare) were washed twice in the previously described liver lysis buffer. 250 µg of liver protein extracts were diluted in 500 µl of lysis buffer containing 1 mM DTT and a cocktail of protease inhibitor and incubated for 2 h on a rotating wheel at 4°C with 20 µl of beads. After incubation, cap-binding-proteins coated beads were washed five times in 500 µl of liver lysis buffer containing 0.5 mM PMSF and 1 mM DTT. 7-methyl GTP bound proteins were eluted by SDS-PAGE loading buffer, separated by SDS-PAGE, transferred to PVDF membranes, and analysed by Western blotting as described.

Mean and standard error of the mean were computed for each time point. The rhythmic characteristics of the expression of each gene or protein were assessed by a Cosinor analysis [74]. This method characterizes a rhythm by the parameters of the fitted cosine function best approximating the data. A period of 24 h was a priori considered. The rhythm characteristics estimated by this linear least squares method include the mesor (rhythm-adjusted mean), the double amplitude (difference between minimum and maximum of fitted cosine function), and the acrophase (time of maximum in fitted cosine function). A rhythm was detected if the null hypothesis was rejected with p<0.05. In such a case, the 95% confidence limits of each parameter were computed. The Cosinor 2.3 software used in this study has been elaborated by the Circadian Rhythm Laboratory at University of South Carolina and is freely available at this address: http://www.circadian.org/softwar.html↗. The statistical significance of differences in the mesor was evaluated by a Student's t-test.