Distinct unfolded protein responses mitigate or mediate effects of nonlethal deprivation of C. elegans sleep in different tissues
- Jarred Sanders†1Email authorView ORCID ID profile,
- Monika Scholz†2,
- Ilaria Merutka2 and
- David Biron1, 2, 3
© Sanders et al. 2017
Received: 5 June 2017
Accepted: 24 July 2017
Published: 28 August 2017
Disrupting sleep during development leads to lasting deficits in chordates and arthropods. To address lasting impacts of sleep deprivation in Caenorhabditis elegans, we established a nonlethal deprivation protocol.
Deprivation triggered protective insulin-like signaling and two unfolded protein responses (UPRs): the mitochondrial (UPRmt) and the endoplasmic reticulum (UPRER) responses. While the latter is known to be triggered by sleep deprivation in rodent and insect brains, the former was not strongly associated with sleep deprivation previously. We show that deprivation results in a feeding defect when the UPRmt is deficient and in UPRER-dependent germ cell apoptosis. In addition, when the UPRER is deficient, deprivation causes excess twitching in vulval muscles, mirroring a trend caused by loss of egg-laying command neurons.
These data show that nonlethal deprivation of C. elegans sleep causes proteotoxic stress. Unless mitigated, distinct types of deprivation-induced proteotoxicity can lead to anatomically and genetically separable lasting defects. The relative importance of different UPRs post-deprivation likely reflects functional, developmental, and genetic differences between the respective tissues and circuits.
Disrupting mammalian sleep during development correlates with negative effects on physical, cognitive, and social health, suggesting that sleep is important for appropriate development [1–3]. Nonlethal sleep deprivation also causes lasting neurological and behavioral deficits in Drosophila melanogaster . However, a mechanistic grasp of why inadequate sleep during development is particularly deleterious is lacking.
Caenorhabditis elegans exhibits developmentally timed sleep during lethargus, a 2- to 3-h-long period at the termination of each larval stage [5–7]. Similar to mammalian sleep, lethargus is characterized by locomotion and feeding quiescence, sensory gating, a typical posture, rebound sleep, and deeply conserved regulation [6, 8–15].
Severe sleep deprivation activates DAF-16/FoxO, the C. elegans Forkhead box O (FoxO) transcription factor. FoxOs function broadly in regulating metabolism, lifespan, and responses to environmental stressors [16–24]. Nuclear translocation of DAF-16/FoxO is inhibited by the insulin/insulin-like growth factor signaling (IIS) pathway [17, 18]. In response to prolonged and continuous sleep deprivation, DAF-16 translocates to the nucleus to mitigate or delay lethality . DAF-16 is also required for rebound sleep following much weaker disruptions .
A particular proteotoxic outcome of sleep deprivation is endoplasmic reticulum (ER) stress. In all species examined, including rodents and flies, expression of the ER chaperone immunoglobulin binding protein (BiP/Grp78) from the heat shock 70 protein family is upregulated upon sleep deprivation [25–30]. The C. elegans homolog of mammalian BiP is HSP-4 [31–33]. BiP/HSP-4 upregulation requires the action of the ribonuclease inositol-requiring protein-1 (IRE-1), a key receptor for sensing misfolded ER proteins [32–37]. IRE-1 signaling activates the XBP-1 transcription factor, which changes the expression of BiP and other genes in the deeply conserved unfolded protein response (UPRER) pathway [25, 28, 30, 38–41].
Prolonged wakefulness increases daily energy expenditure [42–44], for instance, in the brain [45, 46]. Consequently, energy production by the mitochondrial oxidative phosphorylation system is upregulated [25, 30, 47–50]. Nevertheless, the roles of the mitochondrial UPR (UPRmt) following sleep deprivation are largely unknown, although one study found that sleep deprivation induces mitochondrial chaperones (to a lesser degree than BiP) in rat cerebral cortexes .
Upon mitochondrial stress, expression of ubl-5, encoding a ubiquitin-like protein, is upregulated, and UBL-5 plays a key role in activating dedicated chaperones and proteases of the UPRmt [51–54]. In C. elegans, chemically induced mitochondrial stress upregulates the mitochondrion-specific chaperones HSP-6 and HSP-60 (from the Hsp70 and Hsp10/16 superfamilies) [51, 55].
Tractable model organisms have been prominently used to study responses to environmental stressors, such as oxidation or heat [56, 57]. In contrast, C. elegans sleep deprivation was minimally explored, and its lasting impacts, other than lethality, were never characterized. Here we established an automated approach to inflicting severe yet nonlethal deprivation of developmentally timed sleep in C. elegans. We found that worm sleep deprivation inflicts both mitochondrial and ER stress, as indicated by the triggering of the UPRmt and the UPRER. When the UPRs were genetically impeded, lasting defects in feeding, fecundity, and egg-laying physiology were detected. Moreover, different UPRs protected different tissues from the impacts of deprivation.
To assay feeding, we measured the pumping motion of the pharynx, a neuromuscular organ that takes in bacterial food, expels excess liquid, and passes food to the intestine [58, 59]. Maintaining these functions requires speed and regularity and is energetically costly. The pharyngeal nervous system consists of 20 neurons, is isolated from the rest of the animal by the basal lamina, and can operate independently [60, 61]. The rate of pumping depends on feeding history, quality of food, and endogenous serotonin levels [62–65]. In addition, pumping can be stimulated with exogenous serotonin [66, 67]. We found that sleep deprivation-induced mitochondrial stress impacts pharyngeal neurons and slows pumping.
In contrast, sleep deprivation-induced ER stress resulted in germ cell apoptosis and abnormal activity in the egg-laying circuit. The key determinant of C. elegans brood size is the number of available sperm . Germ cell apoptosis can be triggered to protect sperm against DNA damage or environmental stressors that are not directly genotoxic. In both cases, highly conserved core apoptotic genes are strictly required for the initiation of programmed cell death [69–73]. One of these is CED-3, a cysteine-aspartate protease essential for execution of apoptosis [74–79]. Conveniently, this process can be visualized: the transmembrane receptor CED-1 mediates engulfment of early apoptotic corpses by surrounding sheath cells [80, 81]. Thus, the translational ced-1::gfp reporter is used to indicate the occurrence of germ cell apoptosis .
In addition, the egg-laying circuit exhibited a post-deprivation defect similar to the outcome of genetically ablating an egg-laying command neuron . Opposite to the case of the feeding circuit, the UPRER (but not the UPRmt) mitigated the impact of deprivation in the egg-laying circuit. Collectively, these findings implicate two UPRs and insulin/insulin-like growth factor signaling in mitigating the impacts of disrupting worm sleep. They show that developmentally timed sleep is a vulnerable period: external stimuli that are benign outside of lethargus are proteotoxic when administered during lethargus. Adequate sleep promotes normal functions in tissues differing in developmental dynamics and physiological activity, and distinct UPRs mitigate different impacts of nonlethal deprivation.
Substantial deprivation of lethargus quiescence can be automatically inflicted
Manually forced locomotion during worm sleep drives translocation of DAF-16 to nuclei of intestinal and body wall muscle cells . We asked whether our 3-min on/off disruptions would induce similar translocation of DAF-16. To address this, we exposed animals expressing fluorescently labeled DAF-16 to 1 h of the vibration stimuli during the first half of the fourth lethargus stage (L4 lethargus). In agreement with the manual (lethal) deprivation protocol, we observed nuclear localization in intestinal cells (Fig. 1c). Translocation was not observed following a mock perturbation protocol, where vibrations were not applied during an equivalent 1-h period. In our hands, a clear translocation response was not observed in body wall muscles.
In contrast to the consequences of continuous manual deprivation , we did not observe any molting defects or lethality following our deprivation conditions. Possibly, this was a consequence of not depriving the animals of quiescence for a continuous period that exceeded 3 min. Collectively, these data demonstrate the ability to automatically and severely disrupt quiescence to a stressful yet nonlethal degree.
Considering these results, the experiments described throughout the manuscript employ three types of deprivation conditions: 1 h of disruptions for acute responses in individual animals expressing fluorescent markers (“mock” animals were loaded to identical observation chambers), 4 h of disruptions for gene expression assays in small groups of tightly synchronized animals, and 12 h of disruptions for lasting effects of deprivation assayed in large groups of animals. To control for nonspecific effects of the prolonged stimulation period, we compared sleep-deprived animals to those exposed to vibrations outside of lethargus. These groups were labeled “control” (see Methods section for details). In the latter two protocols (4 h and 12 h), sleep was disrupted for no more than the 3-h duration of lethargus. To identify effects of vibrations that are nonspecific to lethargus, we assayed a subset of strains without exposing them to any vibrations. These groups were labeled “unperturbed,” and they consistently exhibited similar phenotypes to those of the “control” animals. Thus, nonspecific effects of vibrations were found to be minor and genetically separable from impacts of sleep deprivation.
Two unfolded protein responses are triggered by nonlethal deprivation of C. elegans sleep
Neither mock deprivation nor 1 h of vibration stimuli prior to lethargus affected hsp-4 expression. In contrast, sleep-deprived animals significantly upregulated the expression of hsp-4 (Fig. 2b). Consistently, hsp-4 expression remained elevated after a 4-h period of disruptions, as indicated by quantitative PCR (Fig 2c). Upregulation of hsp-4 requires essential components of the UPRER including IRE-1 and XBP-1 [33, 34, 87]. Consistent with induction of ER stress by environmental stressors or genetic perturbations, sleep deprivation failed to upregulate hsp-4 expression on ire-1 or xbp-1 mutant backgrounds (Fig. 2d).
We similarly assayed two additional indicators of the UPRmt: the hsp-6 and hsp-60 transcriptional reporters. Expression of hsp-6 was broad and most clearly visible in the intestine. Strong expression of hsp-6 or an accumulation of the reporter led to bright staining of a posterior segment of the intestine. Sleep deprivation did not affect the posterior bright patch, but expression in the rest of the intestine was upregulated after the 1-h deprivation protocol. Mock deprivation did not affect hsp-6 expression (Fig. 3b). Real-time PCR was used to assay the relative expression of these genes after a 4-h period of administering the disruptive stimuli that included L4 lethargus. Consistently, we observed elevated expression of ubl-5 and hsp-6 in deprived animals (Fig. 3c). In our hands, we could not detect upregulation of hsp-60 post-deprivation (Additional file 1: Figure S1). Combined, these results show that nonlethal deprivation of worm sleep is proteotoxic and induces both mitochondrial and ER stress. Comparable stimulation outside of lethargus did not trigger these UPRs, indicating that the period of sleep is particularly vulnerable.
The UPRmt plays a role in mitigating effects of nonlethal sleep deprivation on pumping
Pharyngeal pumping for the purpose of feeding is an energetically demanding behavior that can readily be quantified . To identify lasting effects of sleep deprivation on pumping, each animal was assayed for 60 min at a food concentration corresponding to an optical density (OD600) = 2.5 of the bacterial suspension, where pumping activity was intermediate (Additional file 2: Figure S2). Since control and unperturbed animals (as defined above) exhibited nearly identical feeding behaviors, deprivation conditions were typically compared to stimulation outside of lethargus (control).
To control for potential biomechanical impacts of the vibrations, we assayed worms expressing deg-3(u662), a degeneration-causing constitutively active nicotinic acetylcholine receptor (nAChR) channel subunit, in touch neurons expressing the mec-10 gene [89–93]. These transgenics did not respond to vibrations (Additional file 4: Figure S4). In the absence of mec-10 expressing neurons, vibration stimuli during lethargus did not affect subsequent pumping on both wild-type and daf-16 mutant backgrounds (Fig. 4a and Additional file 3: Figure S3). These data indicate that mechanosensation by mec-10 expressing neurons and the ensuing loss of quiescence are required for pumping fatigue following sleep deprivation.
To test whether the UPRmt plays a role in mitigating consequences of nonlethal sleep deprivation, we examined feeding in ubl-5 mutants. Deprived ubl-5 mutants exhibited a decrease in pumping rate as compared to the control group. This pumping defect was rescued by expressing the Publ-5::ubl-5::gfp translational reporter (Fig. 4b and Additional file 3: Figure S3). In contrast, UPRER defective worms did not exhibit pumping fatigue: animals carrying two ire-1 putative null alleles and hsp-4 mutants did not significantly change their mean pumping rate post-deprivation (Fig. 4b). A mild change in the duty ratio of deprived ire-1 mutants may indicate a weak contribution of the UPRER maintaining post-deprivation pumping (Additional file 3: Figure S3). These results demonstrate that ER proteotoxic stress, unlike mitochondrial stress, does not play a major role in mitigating pumping fatigue following sleep deprivation.
Complementarily, electropharyngeograms (EPGs) enabled us to precisely time contractions and relaxations of the pharyngeal corpus and terminal bulb. We therefore used EPGs to measure durations of individual pumps and to confirm our optical measurements of pumping rates (Additional file 5: Figure S5, Additional file 6: Figure S6, and Additional file 7: Figure S7). We found that repeated mechanical stimuli extended the duration of individual pumps. However, the extension of single pumps was not specific to sleep deprivation; deprived animals and controls exposed to the mechanical stimuli before and after lethargus exhibited similar pump durations. The extended duration of individual pumps in ubl-5 mutants was comparable to that of wild-type animals and daf-16 mutants (Additional file 6: Figure S6). Thus, we could differentiate between deprivation-related and nonspecific impacts of mechanical stimuli, and we did not identify a role for the UPRmt in mitigating the nonspecific effect. Combined, our results indicate that the UPRmt (but not the UPRER) mitigates lasting effects of sleep deprivation on pharyngeal pumping.
Nonlethal sleep deprivation impacts pharyngeal pumping by affecting regulatory neurons
The pharynx is isolated from the rest of the animal and can exhibit pumping tens of minutes after it has been dissected out [60, 61]. Pumping defects induced by sleep deprivation can thus originate from pharyngeal regulatory neurons or pharyngeal muscles. We note that our mechanical stimuli do not noticeably affect the buccal plug, a cap of extracellular material that prevents food from entering the pharynx during lethargus , and they do not induce pumping. Consequently, the stimuli do not activate the pharyngeal muscles during lethargus, and “wear and tear” damage caused by anachronistic muscle activation is unlikely.
An additional serotonin receptor, SER-1/5-HT2, is expressed in pharyngeal muscles  but is not required for rapid pumping, whether induced by food or by 5-HT [64, 95]. Consistently, mutants carrying the putative null allele ser-1(ok345) and treated with 5-HT exhibited high pumping rates and no pumping fatigue (Fig. 5a). The absence and presence of rapid 5-HT-induced pumping in ser-7 and ser-1 mutants, respectively, indicates that 5-HT acts through activating pharyngeal neurons. The ability of deprived daf-16 and ubl-5 mutants to pump rapidly suggests that their deficits, exhibited in the presence of food, are the result of regulation rather than a biomechanical defect. In addition, neuronal rescue of DAF-16 function abolished the post-deprivation phenotype (Fig. 5b). Combined, these data indicate that rapid pumping is mechanically possible even in sleep-deprived mutants upon activation of pharyngeal neurons. Thus, sleep deprivation likely leads to lasting deficits in the neural circuit regulating pumping.
Nonlethal deprivation results in UPRER-dependent reduction in brood size
To address the possibility of a floor effect in the daf-16(mgDf50) animals, we increased the brood size of daf-16(mgDf50) mutants through male mating  and observed a 24% reduction in brood size when both hermaphrodites and males were deprived. Disrupting sleep of either hermaphrodites or males resulted in intermediate phenotypes (Additional file 8: Figure S8A).
Since germ cell apoptosis can be enhanced by pharmacologically or genetically induced ER stress , we hypothesized that the UPRER may mediate an adverse impact of sleep deprivation on fecundity. To test this, we compared ire-1 mutants deficient in the UPRER to wild-type animals, daf-16 mutants, and UPRmt deficient ubl-5 mutants. In contrast to the other genotypes, where deprivation reduced fecundity, brood size was not reduced in the absence of IRE-1. Rather, deprived, control, and unperturbed ire-1 mutants were indistinguishable (Fig. 6e). Similarly, fecundity was impervious to sleep deprivation when ASI neurons, required for UPRER-mediated germ cell apoptosis, were genetically ablated (Additional file 8: Figure S8B) [98, 99]. The wild-type phenotype exhibited by ubl-5 mutants served as a negative control and indicated that the UPRmt did not mediate the impact of deprivation on brood size (Fig. 6f).
To control for potential biomechanical effects of the vibrations, we assayed deg-3(u662) transgenics. Failure to respond to vibrations abolished the reduction in brood size in the presence of the stimuli (Fig. 6h, i). Forced locomotion also did not appreciably increase the number of retained eggs (Additional file 9: Figure S9). Overall, these results suggest that nonlethal sleep deprivation in C. elegans negatively impacts brood size and that the UPRER and IIS play distinct roles in this process. Together with the relevance of the UPRmt to pumping fatigue, these data demonstrate distinct UPRs for mitigating impacts of sleep deprivation in different tissues.
Germ cell apoptosis causes the brood size reduction following nonlethal deprivation
The ced-1::gfp translational reporter is an established indicator of germ cell apoptosis . Animals expressing this reporter were subjected to the 1-h deprivation protocol. Deprivation resulted in a significant increase in ced-1::gfp fluorescence, suggesting that germ cell corpses were actively being disposed of. In contrast, the mock deprivation group, i.e., worms not exposed to vibration stimuli, did not exhibit a change in reporter fluorescence during the equivalent period (Fig. 7b).
Next, we counted fluorescently labeled sperm in self-fertilized deprived and control hermaphrodites as previously described [100, 101]. We observed reductions in sperm count following forced locomotion during L4 lethargus as compared to the control groups in wild-type and the two daf-16 strains (Fig. 7c). The reduced sperm counts were sufficient to explain the corresponding reductions in brood sizes. The reductions in daf-16(mgDf50) and daf-16(mu86) mutants were similar, perhaps due to the variability of the measurement, the 3-day assay having fallen short of revealing the full extent of the brood size deficiency, or further loss of sperm during adulthood. Direct indication of germ cell corpse engulfment mediated by CED-1, the requirement of the caspase CED-3, and the reduced sperm count provide three consistent lines of evidence. Together, they show that sleep deprivation induces germ cell apoptosis, consistent with the role of the UPRER in brood size reduction.
The UPRER (but not the UPRmt) mitigates the effects of nonlethal sleep deprivation on activity in the egg-laying circuit
Opposite to the pharyngeal circuit, the egg-laying circuit was affected by ER but not mitochondrial stress. Neither wild-type animals nor UPRmt deficient ubl-5 mutants exhibited abnormal post-deprivation activity. When UPRER deficient ire-1 mutants were deprived, their mean number of vm twitches increased by 30%, mirroring the trend reported for HSN deficient animals (Fig. 8c).
Physiological activity in the egg-laying circuit is coupled with body posture and locomotion during brief periods around egg-laying events [83, 103, 104]. Therefore, we could not rule out the possibility that differences in locomotion over long timescales may also correlate with activity in the egg-laying circuit. Specifically, we asked whether a potential effect of deprivation on locomotion may indirectly cause the vm twitching phenotype observed in ire-1 mutants. To address this, we assayed the effect of sleep deprivation on mean velocity and measured the correlations between mean velocity and physiological activity (twitches) in the vms.
In all genotypes assayed, the mean velocity did not vary significantly between undisrupted and deprived animals (Additional file 10: Figure S10). While correlations between the mean velocity and vm twitching were found in wild-type, UPRER deficient, and UPRmt deficient worms, they were weakest in ire-1 mutants (Additional file 11: Figure S11). Moreover, the differences between mean or median velocities of deprived and nondeprived worms were insufficient to account for the 30% increase in vm twitching. Thus, excess vm twitching was not an indirect consequence of elevated locomotion activity in our assays.
Interestingly, the correlations between activity in the egg-laying circuit and locomotion were stronger in deprived wild-type animals and ubl-5 mutants as compared to their respective undisrupted groups. This trend was reversed in ire-1 mutants (Additional file 11: Figure S11). This observation may indicate that elevated coordination between distinct behaviors during stress may require secreted proteins, such as neuropeptides, whose function depends on processing in the ER . These results support the conclusion that sleep deprivation causes ER stress which, if not mitigated, impacts the egg-laying circuit. Taken together, our data demonstrate that distinct types of proteotoxic stress caused by nonlethal deprivation of worm sleep adversely affected different circuits.
The cognitive, physiological, and behavioral changes resulting from deprivation of human sleep can be subtle and elude superficial phenotyping. Detecting them requires functional imaging and/or proper design of the task being assayed, a clear definition of the sleep deprivation conditions, and careful measurements. Interpreting such results involves consideration of details such as differences in vulnerability to deprivation between individuals and whether the task was monotonous or complex [106–109]. Similarly, wild-type rodents and insects do not typically exhibit gross defects or substantial damage to brain cells following nonlethal deprivation protocols [4, 110–112]. This may be due to protective responses, activated by sleep deprivation, with the capacity to effectively prevent or repair the resulting damage.
To address the lasting impact of sleep deprivation in C. elegans, we established an experimental method enabling severe reduction in quiescence with no lethality or molting defects. Our periodic stimulus allowed for some quiescent behavior to take place throughout lethargus. Possibly, not forcing locomotion for extended continuous periods was key to avoiding a lethal outcome. In contrast, previous work assessed the impact of total sleep deprivation, i.e., consecutive forced movement for 30 min, which resulted in lethal molting defects. The impact on molting was interpreted to indicate a defect in metabolic regulation during lethargus, as the loss of DAF-16 sensitized the animals to this effect . Lasting defects in surviving animals were not previously assayed.
The complete lack of lethality despite a loss of 50% of quiescence during lethargus suggests that quiescence, in and of itself, is an imperfect measure of the quality and restorative benefits of developmentally timed sleep in C. elegans. If homeostatic compensation can affect the quality of sleep , periodically allowing for rebound could confer greater restoration per unit time as compared to uninterrupted sleep. Testing this hypothesis will be key for understanding lethargus and may promote our understanding of additional quiescent states.
Identifying protective responses to worm sleep deprivation may clarify ancient functions of sleep. Cellular damage and repair can vary in molecular detail and occur at various rates. A comparison between tissues with different functions, developmental states, and metabolic demands may resolve requirements for a rest period due to accumulating damage. We found that deprivation triggers the UPRmt, which protects feeding behavior in sleep-deprived worms. In contrast to the UPRER, the UPRmt was not strongly associated with sleep deprivation previously. This may be partly due to a focus on sleep-related changes in gene expression in the brain [25, 27, 28, 38–40, 113–116]. One study reported that the mitochondrial chaperones Hsp60 and glucose-regulated protein 75 (Grp75, from the Hsp70 superfamily) were upregulated in the cerebral cortex of rats after sleep deprivation, although not as much as BiP .
Broad transcriptional responses to sleep deprivation were reported in mouse livers, lungs, and hearts [117, 118]. These studies demonstrate that the molecular consequences of disrupting sleep are not limited to the brain and that sleep contributes to normal function in a manner that may vary between different tissues or organs. For instance, the pharynx of C. elegans is a highly active organ, required to repeatedly generate powerful contractions. Neurons that regulate pumping may face a distinct cellular metabolic challenge and therefore may particularly benefit from the UPRmt post-deprivation.
Nonlethal sleep deprivation upregulates the expression of BiP in rodents and flies, thus indicating the activation of the UPRER [25, 38–40]. We have shown a similar upregulation of HSP-4/BiP in response to disrupting developmentally timed worm sleep. Furthermore, loss of function of the misfolded ER protein receptor IRE-1 affected both fecundity and the egg-laying circuit post-deprivation. These findings in C. elegans show that the activation of the UPRER is a deeply conserved response to sleep deprivation . It remains to be determined whether the complementing set of phenotypes exhibited by the pharyngeal and the egg-laying circuits, whose function depended on the UPRmt and the UPRER, respectively, is associated with differences in their developmental states, energy expenditure, or additional factors.
Finally, we characterized outcomes of nonlethal sleep deprivation with respect to feeding, fecundity, and egg-laying physiology. Consistent with previous findings, our deprivation protocol triggered translocation of DAF-16/FoxO into intestinal cell nuclei  and implicated it in mitigating lasting adverse effects of deprivation. Three lines of evidence indicated that fecundity was reduced due to germ cell apoptosis: a dependence of the effect on CED-3 and IRE-1, a low sperm count in sleep-deprived worms, and the engulfment of early germ cell corpses by surrounding sheath cells as visualized by the ced-1 reporter [74, 75, 81, 98]. Interestingly, sperm quality was recently shown to be negatively impacted by disruptions to sleep in rodents [96, 97].
Hypotheses explaining the core functions of sleep include the notion that its utility may differ across species . The synaptic homeostasis hypothesis proposes that continuous learning during wakefulness is bound to saturate synaptic connections. Therefore, renormalization of net synaptic strength during sleep is required to restore homeostasis [120, 121]. Other suggestions focus on “wear and tear” in physiologically active neurons during wakefulness due to accumulation of protein fragments, unfolded proteins, or other molecular stressors [112, 122–125]. On a larger scale, metabolite clearance from the brain can increase during mammalian sleep . In addition, disrupting sleep is linked to abnormal glucose metabolism and appetite regulation. These findings suggest that sleep is key to normal metabolic and hormonal processes outside the brain [127, 128].
This work describes multiple pathways by which sleep deprivation can upset cellular proteostasis, likely by creating unfavorable biochemical conditions. Specific characteristics of physiological activity and development could affect the balance between accumulation and relief of allostatic load. To the best of our knowledge, how this balance might scale with different types of metabolic loads has not been systematically studied, let alone connected to sleep. Protection of post-deprivation pharyngeal function by the UPRmt is consistent with the notions that sleep reduces cellular metabolic stress and that highly active organs may invoke distinct responses in this context. Implicating the UPRs in mitigating consequences of worm sleep loss indicates that these responses are deeply conserved.
Wild-type, transgenic, and mutant C. elegans strains were cultivated with OP50 bacteria according to standard protocols at 20 °C. The following strains were used: N2 Bristol (wild type), CF1038 daf-16(mu86), GR1307 daf-16(mgDf50), NQ116 muIs211 [pNL213(ges-1p::GFP::daf-16) + rol-6(su1006)] (gift from D. Raizen), NQ441 daf-16(mgDf50); qnIs45[Pdaf-16:GFP::daf-16; Pmyo-2:mCherry] (gift from D. Raizen at the University of Pennsylvania), SJ4151 zcIs19 [ubl-5p::ubl-5::gfp], SJ4100 zcIs13 [hsp-6p::gfp], SJ4058 zcIs9 [hsp-60p::GFP + lin-15(+)], VC2564 ubl-5(ok3389), SJ4200 zcIs41 [ubl-5p::3xmyc-His tag::ubl-5 + myo-3p::gfp]; SJ4151 zcIs19 [ubl-5p::ubl-5::gfp], NQ128 muEx169[unc-119p::GFP::daf-16 + rol-6(su1006)] (gift from D. Raizen at the University of Pennsylvania), DA184 ser-1(ok345), Is[Pmec-10::deg-3(u662)] (gift from M. Treinin at the Hebrew University of Jerusalem), MT3002 ced-3(n1286), MD701 bcIs39 [lim-7p::ced-1::GFP + lin-15(+)], SJ4005 zcIs4 [hsp-4p::gfp], RE666 ire-1(v33), SJ17 xbp-1(zc12), SJ30 ire-1(zc14) II; zcIs4 V, SJ17 xbp-1(zc12) III; zcIs4 V, RB925 ire-1(ok799), VC1099 hsp-4(gk514)II, UX564 jnSi118[Pcomp-1::GFP::H2B::3’comp-1; Cb-unc-119(+)]; him-5(ok1896) (gift from G. Stanfield at the University of Utah), LX1938 egl-1(n986dm)V; vsIs164 X; lite-1(ce314) X; lin-15(n765ts) X (gift from K. Collins at the University of Miami), LX1918 vsIs164 X; lite-1(ce314) X; lin-15(n765ts) X (gift from K. Collins at the University of Miami), IV205 ueEx122 [str-3::TeTx::GFP; elt-2::sl2GFP] (gift from S. Chalasani at the Salk Institute for Biological Studies).
Severe nonlethal deprivation protocol
Motion and quiescence were identified using the image difference method as described in . To disrupt quiescence during lethargus, mid-L4 animals were transferred to 60-mm nematode growth media (NGM) plates containing 10 mL of medium and seeded with 50 μL of OP50 culture that was grown overnight at 37 °C. Vibrations (1 kHz) were delivered using the mechanical clamps described in . In contrast to the brief and well-spaced disruptions described previously, the stimuli described here were composed of 3 min of vibrations interspersed with 3 min of “quiet” intervals, i.e., a period of 6 min and a duty cycle of 50%.
Synchronization was performed in two steps. Gravid adults were allowed to lay eggs on a fresh plate for 2 h. Of the resulting larvae, mid and early L4s were manually picked for the deprived and control groups, respectively. Manually picking the L4 larvae reduced the variability of the timing of lethargus onset to 2 h.
Assays described throughout the manuscript employ one of three deprivation protocols: (1) fluorescent markers were assayed in individual worms during the first half of lethargus following a 1-h period of 3-min on/off disruptions. Control animals were either not stimulated at all (“mock”) or stimulated for an hour prior to lethargus. (2) Gene expression was assayed in small groups of tightly synchronized animals. Deprived groups were exposed to 3-min on/off disruptions for a period of 4 h that included lethargus, and control groups were exposed to 4 h of vibrations outside of lethargus. (3) Lasting defects were assayed in large groups of animals following a 12-h period of 3-min on/off stimulation. The stimulation period was initiated prior to lethargus and terminated at the young adult stage. Control animals were exposed to vibrations 5 h before lethargus, not stimulated for 5 h that included lethargus, and stimulated again for 5 h after lethargus. In cases when animals that were never stimulated were assayed, they were labeled “unperturbed” to differentiate them from the standard “control” groups.
Optical measurements of pharyngeal pumping
Post-stimulus (see above) young adults were picked into liquid NGM and loaded into a WormSpa microfluidic device [65, 130]. An Escherichia coli OP50 overnight culture, concentration-adjusted in NGM to OD600 = 2.5 (an intermediate food concentration), was flown through the device at a constant rate throughout the assay. After 1 h of acclimation in the device, the animals were imaged for an additional 1 h at a magnification of 10× and 62.5 frames per second using a Basler acA1920-25um complementary metal-oxide-semiconductor (CMOS) camera mounted on Celestron 44104 microscopes . Pumping events were identified using a custom Python script which aligned and subtracted consecutive images and calculated the entropy of the difference image. A pumping event resulted in a characteristic spike in this entropy [65, 132].
Worms were loaded to a NemaMetrix ScreenChip System microfluidic device (NemaMetrix, Inc., Eugene, OR, USA) on a standard dissection microscope and allowed to acclimate for at least 30 min before recording. The device was filled with either bacterial food (OD600 = 2.5) or a solution of 10 mM 5-HT in NGM buffer. Immediately before the onset of the measurement, the output tube was disconnected to reduce electromagnetic noise. The pumping frequency was measured as described in the ScreenChip User Guide, and each recording lasted 5–10 min (measurements that terminated prematurely were discarded).
Imaging of green fluorescent protein (GFP) reporters
The 1-h deprivation protocol consisted of 3-min-long vibration pulses interspersed with 3-min-long pauses, starting during the first 30 min of L4 lethargus and lasting a total of 1 h. Animals expressing a fluorescent reporter were exposed to these mechanical stimuli in artificial dirt microfluidic devices placed inside a standard petri dish. They were imaged in the device immediately before and after the 1-h period of disruption. Imaging was performed at a magnification of 20× (0.5 numerical aperture, NA) using a Nikon Eclipse Ti microscope (Nikon Inc., Melville, NY, USA) and an Andor iXon X3 EMCCD camera (Andor, Belfast, UK). Fluorescence intensity was determined using custom Python scripts.
Calcium imaging in freely behaving animals
To reduce background fluorescence, calcium imaging was performed in artificial dirt microfluidic devices . Animals co-expressing GCaMP5 and mCherry in their vms were mounted in the presence of bacterial food on an epi-fluorescence Nikon Eclipse Ti inverted microscope. Each worm was imaged at a magnification of 20× (0.5 NA) and a frame rate of 6 frames per second. Images were captured with an Andor iXon X3 EMCCD camera. A Dual View (DV2) two-channel system was used for simultaneous imaging of the red and green channels (Photometrix, Tucson, AZ, USA). Each animal was tracked manually and continuously imaged for a total period of 30 min. Calcium transients were analyzed using custom Matlab scripts (The Mathworks Inc., Natick, MA, USA).
Brood size was counted by visual inspection: 10 h after mechanical stimulation ended, animals were transferred to individual 60-mm NGM plates seeded with a 50-μL drop of OP50 (two animals per plate). Plates were scored in the morning and evening of the following 3 days. To avoid the accumulation of hatched offspring, animals were transferred to new plates prior to the evening counts each day. For male mating, two males and two hermaphrodites were transferred to each plate.
Adult Pcomp-1::GFP::H2B::3’comp-1 animals 24 h post-L4 lethargus were sealed into individual “artificial dirt” chambers filled with NGM and 10 μM levamisole. Confocal images of fluorescently labeled sperm were obtained using a Zeiss LSM 880 microscope with a Plan-Apochromat 40×/1.4 oil differential interference contrast (DIC) objective. Image stacks were analyzed using the FARSIGHT Nucleus Editor (http://www.farsight-toolkit.org/wiki/NucleusEditor). We note that when the fluorescent marker was crossed to a daf-16 mutant background, it was not confined to the nuclei. However, individual sperm cells were still detectable. We observed this in all daf-16(mgDf60) mutants and 25% of the daf-16(mu86) mutants.
The lmn-1 gene, encoding the C. elegans nuclear laminin, was chosen as the endogenous control gene. Primers (except for lmn-1, Shaham lab, Rockefeller University, private communication) were designed using Wormbase.org and the National Center for Biotechnology Information (NCBI) Primer-Basic Local Alignment Search Tool (BLAST) software. They were tested for specificity using NCBI BLAST and by agarose gel electrophoresis (using genomic DNA) and purchased from Integrated DNA Technologies (IDT, Coralville, IA, USA). The primers used for lmn-1, hsp-4, ubl-5, and hsp-6 were TCGAGGCGGAAAAGGCTC (Fwd), GCTCCAGCGAGTTCTCTCTC (Rev), GCCGACAAGGAAAAACTCGG (Fwd), GTGGGGTTGGGTTGGGAAA (Rev), ACAAACTGGAACACGATGGGA (Fwd), TCCCTCGTGAATCTCGTAATCC (Rev), AAGAACTCTGGAGGTGACGC (Fwd), and ACGTTGGGGGTTTCTAAAGAT (Rev), respectively. Real-time quantitative PCR amplifications were performed using 25 μL of QuantiTect SYBR Green Real-Time PCR master mix (QIAGEN, Hilden, Germany), 2 μL of diluted reverse transcription product (2 ng/reaction), 1.5 μL each of forward and reverse primer, and 20 μL of DNase/RNase-free water in a total volume of 50 μL. Amplification was carried out in an AB 7900 HT Real-Time PCR cycler (Applied Biosystems, Foster City, CA, USA) with initial polymerase activation at 95 °C for 15 min, followed by cycles of 94 °C for 15 s denaturation, 57 °C for 30 s for primer-specific annealing, and 72 °C for 30 s for extension. A melting curve analysis was carried out (60–95 °C) to verify the specificity of amplicons, i.e., the absence of primer dimers and nonspecific products. Each assay included six technical replicates and a no-template control for every primer pair.
Pairwise comparisons of data represented in bar or box plot were done using the Student’s t test. In the case of multiple comparisons, significance was calculated using a one-way analysis of variance (ANOVA) test and the Bonferroni post hoc correction. Distributions represented by histograms were compared using the k-sample Anderson-Darling test and the Bonferroni post hoc correction for multiple comparisons (when applicable).
For each figure panel, the following p values are listed in order of the positions of asterisks and not significant (n.s.) labels (top to bottom and then left to right). Fig. 1b: 1.1 × 10–3, 6.5 × 10–15, 2.2 × 10–15, 9.5 × 10–7. Fig. 1c: 0.59, 0.84, 0.002, 0.001. Fig. 2b: 0.003, 0.41, 0.63. Fig. 2c: 4.9 × 10–5. Fig. 2d: 0.36, 0.98, 0.033, 0.164. Fig. 3a: 0.016, 0.11, 0.34. Fig. 3b: 0.04, 0.22. Fig. 3c: 0.003, 0.001, 0.005, 0.002. Fig. 4a: 0.21, 0.048, 0.010, 0.50, 0.26, 0.46. Fig. 4b: 0.006, 0.40, 0.142, 0.074, 0.42. Fig. 5a: 0.018, 0.095, 0.38. Fig. 5b: 0.50. Fig. 6a: 4.8 × 10–3 . Fig. 6b: 5.4 × 10–14. Fig. 6c: 6.1 × 10–3. Fig. 6d: 0.39. Fig. 6e: 0.33. Fig. 6f: 0.007. Fig. 6h: 0.19. Fig. 6i: 0.48. Fig. 7a: 0.46. 0.18. Fig. 7b: 0.009, 0.44. Fig. 7c: 0.050, 0.043, 0.009, 0.008. Fig. 8c: 0.38, 0.017, 0.79. Additional file 1: Figure S1: 0.43. Additional file 3: Figure S3A: 0.37, 0.44, 0.003, 0.50, 0.34, 0.41. Additional file 3: Figure S3B: 0.048, 0.028, 0.40, 0.070, 0.27, 0.50. Additional file 5: Figure S5: 0.16, 0.050, 0.010. Additional file 6: Figure S6B: 0.035, 0.042, 0.021. Additional file 6: Figure S6C: 0.002. Additional file 6: Figure S6D: 0.061, 0.005, 0.28. Additional file 7: Figure S7 (OD600 = 2.5): 0.15, 0.37, 0.22, 0.34, 0.092, 0.36, 0.30, 0.30. Additional file 7: Figure S7 (10 mM 5-HT): 0.039, 0.043, 0.22, 0.47, 0.27, 0.30. Additional file 8: Figure S8A: 2.5 × 10–4, 0.82. Additional file 8: Figure S8B: 0.77. Additional file 10: Figure S10: 0.10, 0.34, 0.26, 0.29.
We thank D. Raizen, K. Collins, M. Treinin, G. Stanfield, and S. Chalasani for reagents. Some strains were provided by the Caenorhabditis Genetics Center (CGC), which is funded by the National Institutes of Health (NIH) Office of Research Infrastructure Programs (P40 OD010440). This work was supported by the National Science Foundation (NSF, IOS 1256989 to DB) and the Howard Hughes Medical Institute International (HHMI) Student Research fellowship to MS.
Funding was provided by the NSF (IOS 1256989, to DB).
Availability of data and materials
All data generated or analyzed during this study are included in this published article and its additional files.
JS, MS, and DB conceptualized the study; JS, MS, IM, and DB developed the methodology; JS, MS, and IM were responsible for the investigation; JS, MS, and DB for the software; JS, MS, IM, and DB for the formal Analysis; and JS, MS, and DB for the visualization. DB wrote the original draft, and JS, MS, IM, and DB reviewed and edited the manuscript. DB was responsible for funding acquisition and supervision. All authors read and approved the final manuscript.
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