Introduction: In the hippocampus, clock gene expression is important for memory and mood; however, the signaling mechanism controlling clock gene expression in the hippocampus is unknown. Recent findings suggest that circadian glucocorticoid rhythms driven by the suprachiasmatic nucleus (SCN) control rhythmic clock gene expression in neurons; in addition, dexamethasone modulates hippocampal clock gene expression. We therefore hypothesized that oscillations of clock genes in the hippocampus could be driven by SCN-controlled circadian rhythms in glucocorticoids. Methods: Temporal profiles of hippocampal clock gene expression were established by quantitative reverse-transcription real-time PCR on rat hippocampi, while cellular distribution was established by in situ hybridization. To determine the effect of rhythmic glucocorticoids on hippocampal clock gene expression, the SCN was lesioned, adrenal glands removed and a 24 h exogenous corticosterone rhythm at physiological levels was reestablished by use of a programmable infusion pump. Results: Daily rhythms were detected for Per1, Per2, Bmal1, Nr1d1, and Dbp, while clock gene products were confirmed in both the hippocampus proper and the dentate gyrus. In sham controls, differential hippocampal expression of Per1 and Dbp between ZT3 and ZT15 was detectable. This rhythm was abolished by SCN lesion; however, reestablishing the natural rhythm in corticosterone restored differential rhythmic expression of both Per1 and Dbp. Further, a 6 h phase delay in the corticosterone profile caused a predictable shift in expression of Nr1d1. Conclusion: Our data show that rhythmic corticosterone can drive hippocampal clock gene rhythms suggesting that the SCN regulates the circadian oscillator of the hippocampus by controlling the circadian rhythm in circulating glucocorticoids.

All mammals, including humans, have an internal 24 h biological clock enabling us to synchronize and optimize our physiology to daily life. The 24 h circadian clock of body is localized in neurons of the suprachiasmatic nucleus (SCN) located in the hypothalamus of the brain and is characterized by rhythmic activity of special clock genes. Circadian 24 h rhythms are present throughout the brain, including the hippocampus, where clock genes seem to be important for memory and mood. However, the signaling mechanism controlling circadian clock gene rhythms in the hippocampus is unknown. We hypothesized that oscillations of clock genes in the hippocampus could be driven by a well-established SCN-controlled circadian rhythm in glucocorticoid hormones from the adrenal glands, known as stress hormone or cortisol in humans and corticosterone in rodents. To test this hypothesis, we generated a rat surgical animal model, in which the SCN and adrenal glands were removed and a 24 h exogenous corticosterone rhythm at normal physiological levels was reestablished by use of an implanted programmable infusion pump. Removal of the SCN efficiently abolished clock gene rhythms in the hippocampus; however, reintroducing the natural glucocorticoid rhythm led to reestablishment of 24 h activity rhythms in hippocampal clock genes. Thus, our data suggest that rhythmic hormone release from the adrenal glands drives hippocampal clock gene rhythms and therefore provides a dynamic 24 h hormonal link between the circadian clock of the SCN and circadian function of the hippocampus.

The circadian system ensures physiological adaptation to 24 h rhythmic environmental stimuli across all species [1‒3]. In mammals, the central circadian clock resides in the suprachiasmatic nucleus (SCN) of the hypothalamus and is characterized by 24 h self-sustained molecular oscillations in clock gene expression [4‒6]; however, tissue-specific molecular clock gene-based oscillators have been documented in various brain regions [7, 8] including the hippocampus [9, 10]. The hippocampus is known to be important for learning and memory consolidation [11], functions that are influenced by the circadian system [12, 13]. In addition, disrupted clock gene expression in the hippocampus has been linked to depression [9, 14]. The temporal profiles of clock gene expression have been outlined for individual tissues and generally exhibit a phase delay of several hours as compared to the SCN [8, 15], but local peripheral oscillators still seem to be centrally controlled from the SCN [16‒18]. However, the physiological mechanisms linking circadian dynamics of the central clock of the SCN to peripheral oscillating brain areas, such as the hippocampus, are still not fully understood.

Previous in vitro and in vivo studies have shown that clock gene expression responds to glucocorticoids [19, 20], and an SCN-driven hormonal signaling pathway has been proposed to synchronize circadian timing across tissues [21]. Glucocorticoids, mainly cortisol in humans and corticosterone in rodents, released from the adrenal glands display a strong circadian profile with a peak serum concentration attained to the beginning of the active period [22]. Corticosterone readily penetrates the blood-brain-barrier [23] and although not all brain regions reflect the serum concentrations of corticosterone, the brain overall [24] and the hippocampus specifically follow the circadian corticosterone fluctuations of the serum concentration [25]. The glucocorticoid receptor is widely distributed in the brain with a high concentration in the hippocampus [26‒29]. At the molecular level, a functional glucocorticoid receptor response element has been identified in the promoter of the clock genes Per1 [30] and Per2 [31]. A set of studies combining adrenalectomy and injection of corticosterone in rats have reported glucocorticoid-dependent effects on the expression of clock genes in the prefrontal cortex [32], the bed nucleus of the stria terminalis, and the amygdala [7, 33], whereas timely controlled rhythmic infusion of corticosterone at physiological levels has been shown to drive clock gene rhythms in the cerebellum [34]. Further, hippocampal clock gene expression is influenced by high-dose injections of the synthetic steroid dexamethasone [10]. On the other hand, chronic exposure to glucocorticoids in rats has been shown to disrupt rhythms in hippocampal clock gene transcription accompanied by altered synaptic plasticity and memory impairment [30].

We therefore hypothesized that clock gene oscillations in the hippocampus could be driven by SCN-controlled circadian rhythms in glucocorticoids. To test this hypothesis, we employed a surgical rat model, in which lesion of the SCN and adrenalectomy was combined with rhythmic infusion of corticosterone at physiological levels by use of an implanted programmable pump.

Animals

For surgeries, diurnal series, and RNAscope in situ hybridization, male Sprague Dawley rats (120 g) were acquired from Janvier Laboratories (Saint Berthevin Cedex, France). For surgeries, the rats had a minimum of 1 week to acclimatize to the facility prior to the first procedure and were 7 weeks old at euthanasia. For the diurnal series, rats were 6 weeks old (200–250 g) at euthanasia. For RNAscope in situ hybridization, animals were 6 weeks old (200–250 g) at euthanasia. For radiochemical in situ hybridization, male Sprague Dawley rats (150 g) were acquired from Charles River (Sulzfeld, Germany) and euthanized 3 weeks later at the age of 8 weeks (250–300 g). Animals were kept under a controlled 12 h light/12 h dark schedule with food and water ad libitum. Euthanasia was done by CO2 anesthesia followed by decapitation at specific Zeitgeber times (ZTs). Animal handling and sampling during the dark phase were performed in dim red light. All animal experiments were performed according to the EU directive 2010/63/EU with approvals from the Danish Council for Animal Experiments (authorization number 2017-15-0201-01190) and by the Faculty of Health and Medical Sciences at the University of Copenhagen (P21-190).

Surgical Procedures

For the in vivo model for rhythmic corticosterone effects, rats were subjected to a combination of two surgical sessions. All procedures have previously been published in detail [34]. In short, the first surgery included an electrical-induced lesion of the SCN (SCNx) and implantation of a subcutaneous telemetry transmitter. Stereotaxic lesions were performed with a DC Lesion Making Device 3500 (Ugo Basile, Gemonio, Italy) at 1.3 mA for 90 s with electrode tip coordinates relative to bregma: anteroposterior −0.3 mm, mediolateral 0.0 mm, dorsoventral −9.2 mm (−9.0 mm in shams with no current applied). After 1 week of telemetric monitoring to identify arrhythmic animals, a second surgery which included bilateral adrenalectomy (ADX) and pump implantation was performed; the transmitter was transferred to an intra-abdominal placement through the incision made for adrenalectomy.

Procedures were performed in Sevoflurane anesthesia with analgesia consisting of preoperative Carprofen (5 mg/kg s.c.) and Bupivacaine (2.5 mg/mL, s.c. locally). For the second surgery, Buprenorphine (0.03 mg/kg s.c.) was added to the analgetic preoperative protocol and an antibiotic (Enrofloxacin 10 mg/kg/day) was added to drinking water from 2 days prior to surgery to 2 days after surgery. Adrenalectomized animals received 0.9% saline in their drinking water. Two separate groups of animals underwent the SCN lesion combined with ADX and pump implantation: one group was used exclusively for determining diurnal serum corticosterone profiles by repetitive blood sampling, while the second group was undisturbed from pump implantation to tissue and blood sampling at euthanasia.

Telemetry

Telemetry transmitter (TA-F10, Data Science International, St. Paul, MN, USA) implant data were collected by receivers (RCM-1) placed under individual home cages. Activity and body temperature were logged for 10 s every 10 min with data registered in the Dataquest A.R.T.™ acquisition system. Rhythm analysis was performed using the ImageJ plugin ActogramJ [35]. χ2 analyses of activity and temperature were based on the actograms for the last 5 days before euthanasia, and rhythm robustness was calculated here as relative Qp [35, 36].

Corticosterone Pumps

Programmable iPRECIO SMP-200 pumps (Primetech Corporation, Tokyo, Japan) were used to reestablish daily rhythms in serum corticosterone levels. Corticosterone profile, dose range, and technical considerations have previously been described [34]. For this study, a stepwise delivery was used to mimic the natural corticosterone in nocturnal rodents at the transition from light to dark phase. Pumps were programmed in 24 h repetitive loops to deliver corticosterone at 4 μL/h from ZT6-ZT10, 6 μL/h from ZT10-16, 4 μL/h from ZT16-ZT18 and a maintenance dose of 1 μL/h to avoid clotting in the outlet tubing for the remaining 12 h, ZT18-ZT6 (Fig. 1). The pump reservoir was filled with 12.5 mg/mL corticosterone (Sigma-Aldrich, Steinheim, Germany) dissolved in 10% dimethyl sulfoxide in polyethylene glycol 400. The pump content and program in our cohorts of rats resulted in a corticosterone dose range delivery dependent on weight gain throughout the experiment. For determination of the circadian temporal profile of serum corticosterone, the ZT6-ZT18 differentiated high-dose program delivered an average of 2.6 mg/kg/12 h (2.5–2.8 mg/kg/12 h) based on the body weight on the day of blood sampling. The corresponding ZT6-ZT18 high-dose delivery for the surgical series for hippocampus tissue collection received a corticosterone high-dose average of 2.6 mg/kg/12 h (2.3–3.1 mg/kg/12 h) on the first day of pump implantation and an average high dose of 2.4 mg/kg/12 h (2.1–2.6 mg/kg/12 h) on the last experimental day prior to tissue collection. The low maintenance dose of corticosterone of 12 h of 1 μL/h throughout the pump implantation period in both circadian serum series and hippocampal tissue series were all in the range 0.4–0.6 mg/kg/12 h. All controls were implanted with pumps with the identical program of infusion rate for the release of vehicle (10% dimethyl sulfoxide in polyethylene glycol 400).

Fig. 1.

Daily pump program and corticosterone profile. a 24 h program of the implanted pumps releasing corticosterone. The reservoir of the pump contained 12.5 mg/mL corticosterone. b Corticosterone levels in serum from repetitive blood samples obtained in 4 h intervals (ZT3, ZT7, ZT11, ZT15, ZT19, and ZT23) from rats that had undergone a combined surgical protocol of SCN lesion, adrenalectomy, and implant of a corticosterone-releasing pump (SCNx ADX-cort) determined by ELISA. SCN lesions were confirmed by histology and χ2 analysis of telemetric data of running activity and body temperature (data not shown). n = 6. The dashed line displays previously published data from intact rats [34] for comparison.

Fig. 1.

Daily pump program and corticosterone profile. a 24 h program of the implanted pumps releasing corticosterone. The reservoir of the pump contained 12.5 mg/mL corticosterone. b Corticosterone levels in serum from repetitive blood samples obtained in 4 h intervals (ZT3, ZT7, ZT11, ZT15, ZT19, and ZT23) from rats that had undergone a combined surgical protocol of SCN lesion, adrenalectomy, and implant of a corticosterone-releasing pump (SCNx ADX-cort) determined by ELISA. SCN lesions were confirmed by histology and χ2 analysis of telemetric data of running activity and body temperature (data not shown). n = 6. The dashed line displays previously published data from intact rats [34] for comparison.

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Sample Collection

For 24 h clock gene profiling, brains were removed from five animals per time point in 3 h intervals; the right dorsal hippocampus was removed and frozen on dry ice. Brains for in situ hybridization procedures were collected at ZT15 and frozen on dry ice. To establish the daily serum oscillations from the programmable pump, 100 μL blood samples were taken every 4 h starting at ZT7 from 6 rats with SCNx, ADX, and corticosterone in the pump. Blood was collected in uncoated 1.5 mL centrifuge tubes 6 days after the second surgical session (ADX and pump implantation) by a lancet puncture in the tip of the tail in awake rats; sampling was done within 3 min of initial handling of the rat’s home cage. Collected blood was kept for 30 min at room temperature for clotting and then transferred to wet ice for further processing.

SCNx ADX rats were sacrificed at ZT3 and ZT15 and trunk or heart blood was collected and stored as described above. The brain was removed, the right dorsal hippocampus was isolated, and the remaining brain kept intact; tissue was frozen on dry ice. The brain was placed in a cryostat and divided by a coronal section approximately 3 mm posterior to bregma. The anterior part was used for histological verification of SCN lesion from 14 μm coronal sections spanning the length of the SCN (online suppl. Fig. S1; for all online suppl. material, see https://doi.org/10.1159/000533151). From the posterior part of the brain containing the left hippocampus, 1 mm thick coronal slices were prepared; under a microscope, the cornu ammonis 1 (CA1) and the dentate gyrus were carefully dissected (online suppl. Fig. S2) in a cryostat and stored at −80°C for quantitative reverse-transcription real-time PCR (qRT-PCR).

Serum Analysis

Serum corticosterone concentrations were measured using Detect X® Corticosterone Enzyme Immunoassay Kit (Arbor Assays, Ann Arbor, MI, USA; catalog number K014-H1) in accordance with the manufacturer’s instruction for the 100 μL assay format (standards equal to a sample concentration range of 2–500 ng corticosterone/mL serum). In all experiments, concentrations of corticosterone were within the range of these standards.

Quantitative Reverse-Transcription Real-Time PCR

A detailed protocol for RNA isolation, DNAse treatment, cDNA synthesis, and preparation of plasmids for standard curve generation has been previously published [34]. qRT-PCR was run on a Lightcycler 96 (Roche Diagnostics, Hvidovre, Denmark) with a total reaction volume of 10 μL including 0.5 μm primers (Table 1), Faststart Essential DNA Green Master (Roche Diagnostics, Hvidovre, Denmark), and 0.2 μL cDNA (dorsal right hippocampus) or 0.4 μL cDNA (CA1 and the dentate gyrus).

Table 1.

Sequences of primers used for qRT-PCR and probes used for radiochemical in situ hybridization

TranscriptqRT-PCR primer setsIn situ hybridization probes
NameGenBank accessionPositionForward primer sequence (5′-3′)/reverse primer sequence (5′-3′)PositionProbe sequence (5′-3′)
Bmal1 NM_024362.2 2038–2132 ATT​CCA​GGG​GGA​ACC​AGA/GAA​GGT​GAT​GAC​CCT​CTT​ATC​CT 2357–2322 GAT​CCT​TGG​TCG​TTG​TCT​ATC​ATG​TCG​ATG​CCT​ATG 
Clock NM_021856.3 2402–2483 CAG​CCG​CAT​CCT​TCA​GTT/CAT​GGA​GCA​ACC​GAG​ATG​T   
Per1 NM_001034125.1 2411–2574 ACA​CCC​AGA​AGG​AAG​AGC​AA/GCG​AGA​ACG​CTT​TGC​TTT​AG 699–664 GAG​AGT​GTA​TTC​AGA​TGT​GAT​ATG​CTC​CAA​TTC​CTC 
Per2 NM_031678.2 3319–3409 CAT​CTG​CCA​CCT​CAG​ACT​CA/CTG​GTG​TGA​CTT​GTA​TCA​CTG​CT 320–283 GTG​AGT​GTT​GGA​CGA​TTC​CAC​TAA​CAT​CCG​CAG​CTC​CT 
Cry1 NM_198750.2 2046–2147 CCT​TCT​AAT​CCT​AAT​GGG​AAC​G/ACC​ACT​TCC​TTG​AGA​GCA​GTT​T   
Cry2 NM_133405.2 303–414 GTG​TTC​CCA​AGG​CTT​TTC​AA/CCT​CCT​TGG​CCA​TCT​TCA​TA 954–919 CCG​CTG​TAT​AGA​AGA​ATT​CTC​GCC​ATA​GGA​GTT​GTC 
Nr1d1 NM_001113422.2 1878–2167 GCA​CGA​CCA​GGT​GAC​CCT​GCT/GCT​GCT​CCA​CCG​AAG​CGG​AAT​T   
Dbp NM_012543.3 1131–1301 GCA​AGG​AAA​GTC​CAG​GTG​CCC​G/CTC​CTG​CCG​CAA​TAG​GGC​GTT   
Gapdh NM_017008.4 77–386 TGG​TGA​AGG​TCG​GTG​TGA​ACG​GAT/TCC​ATG​GTG​GTG​AAG​ACG​CCA​GTA   
TranscriptqRT-PCR primer setsIn situ hybridization probes
NameGenBank accessionPositionForward primer sequence (5′-3′)/reverse primer sequence (5′-3′)PositionProbe sequence (5′-3′)
Bmal1 NM_024362.2 2038–2132 ATT​CCA​GGG​GGA​ACC​AGA/GAA​GGT​GAT​GAC​CCT​CTT​ATC​CT 2357–2322 GAT​CCT​TGG​TCG​TTG​TCT​ATC​ATG​TCG​ATG​CCT​ATG 
Clock NM_021856.3 2402–2483 CAG​CCG​CAT​CCT​TCA​GTT/CAT​GGA​GCA​ACC​GAG​ATG​T   
Per1 NM_001034125.1 2411–2574 ACA​CCC​AGA​AGG​AAG​AGC​AA/GCG​AGA​ACG​CTT​TGC​TTT​AG 699–664 GAG​AGT​GTA​TTC​AGA​TGT​GAT​ATG​CTC​CAA​TTC​CTC 
Per2 NM_031678.2 3319–3409 CAT​CTG​CCA​CCT​CAG​ACT​CA/CTG​GTG​TGA​CTT​GTA​TCA​CTG​CT 320–283 GTG​AGT​GTT​GGA​CGA​TTC​CAC​TAA​CAT​CCG​CAG​CTC​CT 
Cry1 NM_198750.2 2046–2147 CCT​TCT​AAT​CCT​AAT​GGG​AAC​G/ACC​ACT​TCC​TTG​AGA​GCA​GTT​T   
Cry2 NM_133405.2 303–414 GTG​TTC​CCA​AGG​CTT​TTC​AA/CCT​CCT​TGG​CCA​TCT​TCA​TA 954–919 CCG​CTG​TAT​AGA​AGA​ATT​CTC​GCC​ATA​GGA​GTT​GTC 
Nr1d1 NM_001113422.2 1878–2167 GCA​CGA​CCA​GGT​GAC​CCT​GCT/GCT​GCT​CCA​CCG​AAG​CGG​AAT​T   
Dbp NM_012543.3 1131–1301 GCA​AGG​AAA​GTC​CAG​GTG​CCC​G/CTC​CTG​CCG​CAA​TAG​GGC​GTT   
Gapdh NM_017008.4 77–386 TGG​TGA​AGG​TCG​GTG​TGA​ACG​GAT/TCC​ATG​GTG​GTG​AAG​ACG​CCA​GTA   

Radiochemical in situ Hybridization

Radiochemical in situ hybridization was performed on 12 μm cryostat sections by using 35S-labeled DNA oligo probes (Table 1) as previously described [37].

RNAscope in situ Hybridization

RNAscope in situ hybridization was performed on 12 μm cryostat sections as previously described [38] following the protocol for fresh frozen sections (ACDBio; #320513, #320293, and #320850) by use of probe sets detecting the glucocorticoid receptor Nr3c1 (positions 408–1,401 on NM_012576.2, C1-tagged), Per1 (positions 139–1,112 on NM_001034125.1, C3-tagged), and Per2 (positions 3,647–4,603 on NM_031678.1, C3-tagged), respectively.

Statistical Analysis

Data are presented as individual data points with means (and standard error of mean in bar graphs) for each group. Data were analyzed with the JTK-Cycle script in R software for the 24 h time series [39] and two-way ANOVA for surgical experiments followed by Bonferroni post hoc tests in GraphPad Prism Version 9.4.1 (GraphPad Software, San Diego, CA, USA). n values are given in the figure legends. For telemetric analyses, the relative Qp values for both locomotor activity and body temperature for each animal were calculated by dividing the Qp value defined by the ActogramJ χ2 analysis at 1,440 min (24 h) by the total number of data registrations in the 5-day period monitored. In all analyses, a p value of 0.05 was considered to represent statistical significance.

Temporal Clock Gene Expression Profiles in the Rat Hippocampus Reveal Antiphasic Per and Bmal1 Expression

As a first step toward analyzing the potential regulatory role of glucocorticoids in hippocampal clock gene expression, daily expression patterns of clock genes were determined by qRT-PCR analyses of rat hippocampi sampled in 3 h intervals (Fig. 2; Table 2). Daily rhythms in transcript levels were detected for six out of eight clock genes. Among these, Per1 and Per2 had their acrophase in the early night at ZT15 (p < 0.0001 for Per1 and p = 0.0060 for Per2, JTK-Cycle analysis), while Bmal1 expression was in antiphase peaking at ZT3 (p = 0.0028, JTK-Cycle analysis). Nr1d1 and Dbp were rhythmic with peaks late in the light period at ZT9 and ZT10.5, respectively (p values <0.0001, JTK-Cycle analysis). Cry2 was also rhythmic with a peak at ZT10.5 (p = 0.011, JTK-Cycle analysis). Differential daily expression of Cry1 and Clock was not detected. Expanding on previous analyses of clock gene expression in the rat hippocampus [10], radiochemical in situ hybridization showed expression of Per1, Per2, Cry2, and Bmal1 in the pyramidal layer of the cornu ammonis and the granular layer of the dentate gyrus (Fig. 2). Thus, clock genes are expressed throughout pyramidal and granular layers of the hippocampus with Per1 and Per2 oscillating in antiphase to Bmal1.

Fig. 2.

Diurnal clock gene expression profiles in the hippocampus. a qRT-PCR analyses of clock gene expression in rat hippocampi sampled in 3 h intervals. Statistical analyses were performed in JTK-Cycle. n = 5. *, p < 0.05; **, p < 0.01; ****, p < 0.0001. See Table 2 for details on exact p values and estimated peak expression time points. b Representative autoradiographs of radiochemical in situ hybridization on coronal rat brain sections for detection of transcripts encoded by the clock genes Per1, Per2, Cry2, and Bmal1. Animals were sacrificed at ZT15. Clock gene products are detectable in both the cornu ammonis and the dentate gyrus. Scale bar, 1 mm.

Fig. 2.

Diurnal clock gene expression profiles in the hippocampus. a qRT-PCR analyses of clock gene expression in rat hippocampi sampled in 3 h intervals. Statistical analyses were performed in JTK-Cycle. n = 5. *, p < 0.05; **, p < 0.01; ****, p < 0.0001. See Table 2 for details on exact p values and estimated peak expression time points. b Representative autoradiographs of radiochemical in situ hybridization on coronal rat brain sections for detection of transcripts encoded by the clock genes Per1, Per2, Cry2, and Bmal1. Animals were sacrificed at ZT15. Clock gene products are detectable in both the cornu ammonis and the dentate gyrus. Scale bar, 1 mm.

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Table 2.

JTK-Cycle analysis of daily profiles of clock gene expression in the hippocampus

TranscriptBenjamini-Hochberg q valueAdjusted p valuePhase (estimated ZT for peak expression)
Per1 0.000011 0.0000040**** 15 
Per2 0.0096 0.0060** 15 
Cry1 1 n.s 
Cry2 0.015 0.011* 10.5 
Bmal1 0.0056 0.0028** 
Clock 1 n.s 
Nr1d1 0.0000000029 0.00000000070**** 
Dbp 0.0000000014 0.00000000018**** 10.5 
TranscriptBenjamini-Hochberg q valueAdjusted p valuePhase (estimated ZT for peak expression)
Per1 0.000011 0.0000040**** 15 
Per2 0.0096 0.0060** 15 
Cry1 1 n.s 
Cry2 0.015 0.011* 10.5 
Bmal1 0.0056 0.0028** 
Clock 1 n.s 
Nr1d1 0.0000000029 0.00000000070**** 
Dbp 0.0000000014 0.00000000018**** 10.5 

*, p < 0.05; **, p < 0.01; ****, p < 0.0001; n.s., not significant.

Exogenous Corticosterone Infused via Programmable Pumps Can Mimic the Natural Daily Endogenous Serum Profile in Otherwise Arrhythmic Rats

To determine if programmed pumps (Fig. 1a) could generate a natural rhythm in serum corticosterone, the diurnal profile of serum corticosterone from animals that had undergone SCN-lesion, adrenalectomy, and implantation with a pump releasing corticosterone (SCNx ADX-cort) was established (Fig. 1b). The serum profiles display a clear peak in corticosterone at the transition from light to dark with the highest values registered in samples from ZT11 and ZT15 followed by a subsequent decline with lowest values registered at the night-day transition corresponding to the profile in intact rats (Fig. 1b).

To verify the corticosterone rhythms and general diurnal profiles of physiological parameters of the surgical groups included in investigations of the regulation of clock gene expression in the hippocampus, all animals had blood collected at euthanasia at ZT3 and ZT15, respectively, and the daily rhythms in running activity and body temperature were concurrently monitored telemetrically (Fig. 3). In the sham group with a vehicle-filled pump (Sham-veh), a significant day-night variation in corticosterone with low levels at ZT3 and high levels at ZT15 was detectable (p = 0.0064, Bonferroni post hoc test following two-way ANOVA) (Fig. 3a). In SCN-lesioned animals with adrenal glands removed and vehicle in the pump (SCNx ADX-veh), very low levels of corticosterone were detected both at ZT3 and ZT15 with no difference between time points (p > 0.9999, Bonferroni post hoc test following two-way ANOVA), whereas a highly significant difference between corticosterone levels at ZT3 and ZT15 was established in SCN-lesioned rats with adrenalectomy and corticosterone in the pump (SCNx ADX-cort) (p = 0.0062, Bonferroni post hoc test following two-way ANOVA) (Fig. 3a).

Fig. 3.

Characterization of serum corticosterone, activity, and temperature rhythms in experimental groups. a Serum corticosterone from blood sampled at ZT3 and ZT15, respectively, from rats in a sham group with a vehicle-filled pump (Sham-veh), SCN-lesioned animals with adrenal glands removed and vehicle in the pump (SCNx ADX-veh), and SCN-lesioned rats with adrenalectomy and corticosterone in the pump (SCNx ADX-cort). Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. **, p < 0.01. b Rhythm robustness of animals in the three experimental groups. The relative Qp value was obtained by χ2 analyses performed on telemetric data of running activity and body temperature from a 5 day period. The dashed line indicates a the relative Qp value corresponding to a significance level of p = 0.05 calculated by χ2 analysis at a period of 24 h.

Fig. 3.

Characterization of serum corticosterone, activity, and temperature rhythms in experimental groups. a Serum corticosterone from blood sampled at ZT3 and ZT15, respectively, from rats in a sham group with a vehicle-filled pump (Sham-veh), SCN-lesioned animals with adrenal glands removed and vehicle in the pump (SCNx ADX-veh), and SCN-lesioned rats with adrenalectomy and corticosterone in the pump (SCNx ADX-cort). Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. **, p < 0.01. b Rhythm robustness of animals in the three experimental groups. The relative Qp value was obtained by χ2 analyses performed on telemetric data of running activity and body temperature from a 5 day period. The dashed line indicates a the relative Qp value corresponding to a significance level of p = 0.05 calculated by χ2 analysis at a period of 24 h.

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Daily rhythms in running activity and body temperature were monitored by telemetric registrations of all rats subjected to surgeries (Fig. 3b). Determining of rhythm robustness by χ2 analyses of telemetric data from individual rats confirmed that all animals included in the Sham-veh group expectedly displayed daily rhythms in both running activity and body temperature, whereas animals included in SCNx ADX-veh and SCNx ADX-cort groups showed no 24 h rhythmicity in neither activity nor temperature (Fig. 3b). In summary, we have generated an otherwise arrhythmic rat model with a daily rhythm in corticosterone within the normal physiological range.

Rhythmic Infusion of Corticosterone Induces Rhythmic Expression of Per1 and Dbp in the Hippocampus of SCN-Lesioned Animals

To determine the possible effects of rhythmic corticosterone on clock gene expression in the hippocampus, hippocampal samples from animals in the three surgical groups, Sham-veh, SCNx ADX-veh, and SCNx ADX-cort, obtained at ZT3 and ZT15, respectively, were subjected to qRT-PCR; analyses were performed on the entire dorsal hippocampus of the right hemisphere (Fig. 4) and on dissected parts of the dorsal hippocampus, namely CA1 and the dentate gyrus, in the left hemisphere (Fig. 5).

Fig. 4.

Clock gene expression in the hippocampus of SCN-lesioned, adrenalectomized, and rhythmic corticosterone-supplemented animals. qRT-PCR analyses of the expression of clock genes in the dorsal hippocampus from rats in a sham group with a vehicle-filled pump (Sham-veh), SCN-lesioned and adrenalectomized rats with vehicle in the pump (SCNx ADX-veh), and SCN-lesioned and adrenalectomized rats with corticosterone in the pump (SCNx ADX-cort). Animals were sacrificed at ZT3 and ZT15. Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. ****, p < 0.0001; *, p < 0.05.

Fig. 4.

Clock gene expression in the hippocampus of SCN-lesioned, adrenalectomized, and rhythmic corticosterone-supplemented animals. qRT-PCR analyses of the expression of clock genes in the dorsal hippocampus from rats in a sham group with a vehicle-filled pump (Sham-veh), SCN-lesioned and adrenalectomized rats with vehicle in the pump (SCNx ADX-veh), and SCN-lesioned and adrenalectomized rats with corticosterone in the pump (SCNx ADX-cort). Animals were sacrificed at ZT3 and ZT15. Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. ****, p < 0.0001; *, p < 0.05.

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Fig. 5.

Clock gene expression in subregions of the hippocampus in response to rhythmic infusion of corticosterone. qRT-PCR analyses of clock gene expression in the dentate gyrus (DG) and CA1 of rats in a sham group with vehicle in the pump (Sham-veh), SCN-lesioned and adrenalectomized rats with vehicle in the pump (SCNx ADX-veh), and SCN-lesioned and adrenalectomized rats with corticosterone in the pump (SCNx ADX-cort). Animals were sacrificed at ZT3 and ZT15. Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. ****, p < 0.0001; **, p < 0.01; *, p < 0.05.

Fig. 5.

Clock gene expression in subregions of the hippocampus in response to rhythmic infusion of corticosterone. qRT-PCR analyses of clock gene expression in the dentate gyrus (DG) and CA1 of rats in a sham group with vehicle in the pump (Sham-veh), SCN-lesioned and adrenalectomized rats with vehicle in the pump (SCNx ADX-veh), and SCN-lesioned and adrenalectomized rats with corticosterone in the pump (SCNx ADX-cort). Animals were sacrificed at ZT3 and ZT15. Statistical analyses were performed by two-way ANOVA followed by Bonferroni post hoc tests. n = 3–6. ****, p < 0.0001; **, p < 0.01; *, p < 0.05.

Close modal

In the right dorsal hippocampus (Fig. 4), Sham-veh rats exhibited significant differential expression of Per1 and Dbp between ZT3 and ZT15 (p values <0.0001, Bonferroni post hoc tests following two-way ANOVA). These rhythms disappeared in the SCNx ADX-veh rats but were reestablished by rhythmic corticosterone infusion in SCNx ADX-cort animals (p = 0.010 for Per1 and p = 0.031 for Dbp, Bonferroni post hoc tests following two-way ANOVA). Per2, Cry1, Bmal1, and Nr1d1 were not differentially expressed between ZT3 and ZT15 in neither the Sham-veh group nor in the SCNx ADX-veh animals; however, rhythmic infusion of corticosterone generated a rhythm in Nr1d1 in the SCNx ADX-cort group with the highest levels detected at ZT3 (p = 0.011, Bonferroni post hoc test following two-way ANOVA).

In dissected subregions of the hippocampus of sham-veh rats, namely CA1 and the dentate gyrus (Fig. 5), Per1 and Dbp were found to retain the same magnitude of change from ZT3 to ZT15 (p values <0.0001, Bonferroni post hoc tests following two-way ANOVA) as in the whole dorsal hippocampus. Also, rhythms in both genes disappeared in the SCNx ADX-veh group. In case of Per1, rhythmic corticosterone in SCNx ADX-cort rats reestablished a rhythm in both CA1 and the dentate gyrus (p values <0.0001, Bonferroni post hoc tests following two-way ANOVA), whereas for Dbp, significant corticosterone-induced differential expression between day and night was limited to the dentate gyrus (p = 0.013, Bonferroni post hoc test following two-way ANOVA). In the Sham-veh animals, Cry1 expression did not oscillate between ZT3 and ZT15 in neither the dentate gyrus nor CA1. On the other hand, a difference between day and night in Per2 expression was detected in both hippocampal subregions with highest transcript levels at ZT15 (p = 0.019 for CA1 and p = 0.023 for the dentate gyrus, Bonferroni post hoc tests following two-way ANOVA), whereas Bmal1 was also found to be differentially expressed in sham-veh rats with highest levels obtained at ZT3 in both CA1 (p < 0.0001, Bonferroni post hoc test following two-way ANOVA) and in the dentate gyrus (p = 0.014, Bonferroni post hoc test following two-way ANOVA). In both cases, rhythms were abolished in the SCNx ADX-veh rats without being reestablished in the SCNx ADX-cort animals. In agreement with data from the whole dorsal hippocampus, oscillating corticosterone induced a rhythm in Nr1d1 expression in both the dentate gyrus (p = 0.0053, Bonferroni post hoc test following two-way ANOVA) and CA1 (p = 0.031, Bonferroni post hoc test following two-way ANOVA). The overall results of the clock gene differential expression analyses of CA1 and the dentate gyrus resemble the results from the contralateral entire dorsal hippocampus, although some gene-specific distinct regional features of each of the two hippocampal regions are apparent.

To establish if an altered timing of the corticosterone profile in itself would be able to control hippocampal clock gene expression, we employed a previously published experimental protocol [34] resulting in a 6 h phase delay in the corticosterone profile (Fig. 6). A significant difference in hippocampal Nr1d1 expression between ZT3 and ZT15 with high levels at daytime was detected in the sham animals (p = 0.030, Bonferroni post hoc test following two-way ANOVA). Predictably, this rhythm disappeared in the SCN-lesioned animals. Rhythmic infusion of corticosterone with a 6 h delay in SCN-lesioned animals resulted in a significant difference between transcript levels at ZT3 and ZT15 (p = 0.016, Bonferroni post hoc test following two-way ANOVA), but in this group, the Nr1d1 transcript rhythm was inverted with high levels during nighttime. Thus, in this experimental setting, rhythmic expression of Nr1d1 was found to be inverted in the hippocampus corresponding to a predicted 6 h delay in the diurnal clock gene profile, suggesting that hippocampal expression of Nr1d1 follows the corticosterone profile in an otherwise arrhythmic animal.

Fig. 6.

Nr1d1clock gene expression in the hippocampus of rats exposed to a 6 h phase delay in the diurnal corticosterone profile. In the current study, the increasing infusion rate of corticosterone was initiated at ZT6, which will closely mimic the endogenous corticosterone profile. We have previously used the pumps with a rhythmic infusion program; however, the high infusion rate lasting for 12 h was initiated at ZT12 corresponding to a 6 h phase delay [34]. Using this phase delay protocol, we here wanted to test if different timing of the corticosterone oscillation would affect rhythmic clock gene expression profiles accordingly. qRT-PCR analysis for quantification of the Nr1d1 transcript was performed on the dorsal hippocampus of sham animals with vehicle in the pump (Sham-veh), SCN-lesioned animals with vehicle in the pump (SCNx-veh), and SCN-lesioned animals with corticosterone in the pump (SCNx-cort 6 h delay). The pump setting corresponding to a 6 h delay in corticosterone infusion was identical for all groups. The graph on the right-hand side shows the natural rhythm of hippocampal Nr1d1 expression presented in Fig. 2 with a solid line, whereas the dashed line depicts the rhythm artificially delayed by 6 h resulting in opposite rhythms when comparing ZT3 and ZT15. Thus, the inverted rhythm detected in the SCN-lesioned animals exposed to a 6 h phase delay in corticosterone infusion corresponds to a 6 h delay in the natural hippocampal rhythmic profile of Nr1d1.

Fig. 6.

Nr1d1clock gene expression in the hippocampus of rats exposed to a 6 h phase delay in the diurnal corticosterone profile. In the current study, the increasing infusion rate of corticosterone was initiated at ZT6, which will closely mimic the endogenous corticosterone profile. We have previously used the pumps with a rhythmic infusion program; however, the high infusion rate lasting for 12 h was initiated at ZT12 corresponding to a 6 h phase delay [34]. Using this phase delay protocol, we here wanted to test if different timing of the corticosterone oscillation would affect rhythmic clock gene expression profiles accordingly. qRT-PCR analysis for quantification of the Nr1d1 transcript was performed on the dorsal hippocampus of sham animals with vehicle in the pump (Sham-veh), SCN-lesioned animals with vehicle in the pump (SCNx-veh), and SCN-lesioned animals with corticosterone in the pump (SCNx-cort 6 h delay). The pump setting corresponding to a 6 h delay in corticosterone infusion was identical for all groups. The graph on the right-hand side shows the natural rhythm of hippocampal Nr1d1 expression presented in Fig. 2 with a solid line, whereas the dashed line depicts the rhythm artificially delayed by 6 h resulting in opposite rhythms when comparing ZT3 and ZT15. Thus, the inverted rhythm detected in the SCN-lesioned animals exposed to a 6 h phase delay in corticosterone infusion corresponds to a 6 h delay in the natural hippocampal rhythmic profile of Nr1d1.

Close modal

Clock Gene Products Colocalize with the Glucocorticoid Receptor in Cells of the Hippocampus

The glucocorticoid receptor is present in the rat hippocampus [26, 40], but to determine if the receptor is expressed in the clock-containing cells of the hippocampus, RNAscope in situ hybridization was performed (Fig. 7). Cellular colocalization of Per1 mRNA and the glucocorticoid receptor Nr3c1 was detected in both the cornu ammonis (Fig. 7a) and the dentate gyrus (Fig. 7b), and similarly, Per2 and Nr3c1 transcripts colocalized in the cornu ammonis (Fig. 7c) and the dentate gyrus (Fig. 7d), suggesting that corticosterone has the capacity to act directly on gene expression in the clock-containing hippocampal cells.

Fig. 7.

Colocalization of clock gene and glucocorticoid receptor transcripts in neurons of the hippocampus. RNAscope in situ hybridization for colocalization of Per1 and the glucocorticoid receptor Nr3c1 transcripts in CA1 (a) and in the dentate gyrus (b), as well as Per2 and the glucocorticoid receptor Nr3c1 transcripts in CA1 (c) and in the dentate gyrus (d). In each panel, the arrows point to the same cell and the arrow heads point to the same cell photographed in different filter settings. DG, dentate gyrus. Scale bar, 50 μm.

Fig. 7.

Colocalization of clock gene and glucocorticoid receptor transcripts in neurons of the hippocampus. RNAscope in situ hybridization for colocalization of Per1 and the glucocorticoid receptor Nr3c1 transcripts in CA1 (a) and in the dentate gyrus (b), as well as Per2 and the glucocorticoid receptor Nr3c1 transcripts in CA1 (c) and in the dentate gyrus (d). In each panel, the arrows point to the same cell and the arrow heads point to the same cell photographed in different filter settings. DG, dentate gyrus. Scale bar, 50 μm.

Close modal

Circadian oscillators are present throughout the mammalian brain, including the hippocampus as evidenced by 24 h rhythmic clock gene expression in this brain region. The current study was undertaken to identify a physiological signal that would enable the central clock of the SCN to regulate and synchronize extrahypothalamic clock gene rhythms in the hippocampus, and we here by use of an advanced surgical rat model show that rhythmic administration of corticosterone at physiological levels can reintroduce hippocampal clock gene rhythms in otherwise arrhythmic SCN-lesioned animals.

The 24 h profiles of clock gene expression in the rat hippocampus outline the presence of a molecular circadian oscillator in the hippocampus. Previous studies in mice have shown an apparent discrepancy between reported rhythms in different strains within the same species [9, 41]. Although the profiles of a limited number of clock genes have been reported in the rat hippocampus [10, 42], these also seem to differ between strains. Data from our experimental model strain, the Sprague Dawley rat, show that in the positive loop of the transcriptional-translational molecular clockwork, Bmal1 peaks at early morning, while Clock is arrhythmic. In the negative loop, the rhythmic expression of Per1 and Per2 peaks early in the night while their dimer-forming partners Cry1 and Cry2 are either arrhythmic or exhibit a very limited amplitude. The profiles of Nr1d1 and the clock-controlled Dbp are consistent with mouse data [9]. In line with functional circadian clocks in other tissues, Per gene expression is in antiphase to that of Bmal1.

Our finding that hippocampal expression of Per1, Per2, Cry1, and Bmal1 is detectable in the pyramidal layer of the cornu ammonis and the granular layer of the dentate gyrus is consistent with and complements previous reports on the 24 h expression profiles that constitute the clock [10, 42, 43], but notably, our data do not exclude the expression of clock genes in hippocampal glial cells. We expand current knowledge of the glucocorticoid receptor, which has been known to be present in the rat hippocampus for decades [26, 40], to include cellular colocalization with the local hippocampal molecular clock. This colocalization of clock gene transcripts, i.e., Per1 and Per2, and the glucocorticoid receptor shows, in line with the concept outlined below, that circulating glucocorticoids can act directly on clock-containing hippocampal cells.

The synchronizing effect of glucocorticoids on clock gene expression was originally observed when adding serum naturally containing corticosterone to cell cultures [44]. Clock genes rhythmically expressed in extrahypothalamic tissues generally seem to be subject to an overall control by the central clock located in the SCN receiving time-cues to entrain peripheral oscillations; however, in the majority of these peripheral clock gene-expressing brain areas or peripheral tissues, neural connections cannot be traced directly to the SCN. The findings of this report suggest that corticosterone, which exhibits an SCN-controlled circadian rhythm in serum [21], across all regions of the hippocampus, represents a time-of-day signaling mechanism connecting the SCN and hippocampal clock gene expression. That glucocorticoids in this context act directly in the hippocampus is supported by previous reports identifying glucocorticoid response elements in clock gene promoters [31, 45‒47] and the abovementioned cellular colocalization of the glucocorticoid receptor and clock gene transcripts.

As a novel approach, we here determined the effects of rhythmic glucocorticoids on clock gene expression by replicating the exact daily oscillations of endogenous corticosterone with a focus on the normal physiological concentration range. Previous studies have investigated the effect of glucocorticoids by use of the synthetic glucocorticoid agonist dexamethasone [10, 48]; however, considerable differences exist between endogenous corticosterone and dexamethasone. The hippocampus contains glucocorticoid receptors, but also mineralocorticoid receptors [29], and while effect of dexamethasone is primarily mediated through glucocorticoid receptors with only limited effect on the mineralocorticoid receptor [49], corticosterone will bind both types of receptors. The two receptor types bind the same downstream response elements, but studies in rats detect a high occupancy of the mineralocorticoid receptor even at circadian trough levels due to a strong binding affinity of corticosterone [47, 50, 51]. However, even with a high basic occupancy, the amount of mineralocorticoid receptor bound to the glucocorticoid response element in the Per1 promoter still exhibits a circadian rhythm [47] as does the binding of the glucocorticoid receptor [30], indicating that under physiological conditions, both receptor types are susceptible to corticosterone activation during the daily hormone increase, thus highlighting the biological importance of the endogenous ligand. The receptor binding patterns of corticosterone and dexamethasone also vary between brain regions, so while corticosterone is bound in higher concentrations in the hippocampus as compared to the anterior pituitary, the opposite pattern was the case for dexamethasone [52, 53]. In addition, dexamethasone has a biological half-life of 36–72 h and a very different potency profile [54], again supporting the use of corticosterone in experimental designs mimicking normal 24 h physiology.

In the current study, our corticosterone administration procedure was also optimized as compared to previous approaches. Administration of corticosterone by injection in rats would only render a short rise in blood levels, and although a pulse would be sufficient to induce hippocampal Per1 expression [55], it does not mimic the circadian 24 h physiological profile. Studies in mice also show that an injection of corticosterone induces a fast rise in hippocampal hormone concentrations, but at the same time, handling procedures or saline injections would also affect the corticosterone concentration [56], presumably due to a stress response induced by the procedures themselves [22]. Taking advantage of the rat’s nocturnal behavior including an increased consumption of water at night, corticosterone supplementation in the drinking water has also been used to generate a daily rhythm in corticosterone in adrenalectomized animals to analyze the role of glucocorticoids in hippocampal PER2 expression [33]; however, this approach would depend on an intact SCN. As compared to previous approaches, our gain-of-function model based on programmable corticosterone-containing pumps to rescue the hormone rhythm in SCN-lesioned adrenalectomized animals therefore represents normal undisturbed physiology regarding corticosterone serum concentrations and circadian 24 h profile, as well as the natural ligand with its pharmacodynamics and -kinetics.

The surgical model allowed us to isolate the effect of the circadian rhythm in corticosterone from other endogenous circadian signals. In the sham group, expression levels and rhythms generally resembled the diurnal clock gene profiles discussed above; however, oscillations were more distinct in the dissected subregions, suggestively reflecting less biological variation within individual parts of the hippocampus. The SCN lesion and adrenalectomy successfully eliminated diurnal variation in all transcripts in both the whole tissue and subregional analyses. While our general experimental approach does not allow separation of the effect of the aforementioned surgeries, SCN-dependent control of hippocampal clock gene rhythms has previously been described at the protein level for PER2 [7]. With regard to eliminating the source of endogenous corticosterone, some studies found no effect of adrenalectomy on the PER2 protein rhythm in the dentate gyrus [7, 33], while abolishment of hippocampal rhythms in numerous clock genes, including Per2, has also been reported [10], thus preventing clear conclusions on the effect of adrenalectomy on the molecular clockwork of the hippocampus.

At the time points sampled in the current study, rhythmic infusion of corticosterone induced differential expression of Per1, Nr1d1, and Dbp in both whole hippocampi and in dissected dentate gyri and CA1 regions. The effect of rhythmic corticosterone on hippocampal Per1 expression is in accord with our previous study on the corticosterone-induced clock gene rhythms in the cerebellum [34] and also the effect of ultradian pulses of corticosterone on hippocampal Per1 [55]. Interestingly, large-scale analyses of genomic glucocorticoid responses in cell lines determined a distinct sensitivity of Per1 responding to a far lower concentrations than other genes [45, 46]. Combined, these findings suggest that corticosterone could synchronize circadian clocks across tissues by using Per1 as an entrance to the molecular circadian clock. In contrast, a recent study using dexamethasone reported an effect only in the dentate gyrus [10]; this apparent discrepancy probably reflects the above discussed pharmacological differences between the synthetic and the natural agonist and possibly drug administration procedures.

This study expands the group of rhythmic corticosterone-responsive genes in the hippocampus to include the clock-controlled Dbp. At the same time, our experimental design with two sampling time points may prevent detection of corticosterone-induced rhythmicity or phase-shifts in otherwise rhythmic clock genes, such as Per2 and Bmal1. If the whole molecular clock-work of the hippocampus is reactivated or resynchronized by corticosterone, all clock gene rhythms should theoretically be reestablished following the introduction of rhythmic corticosterone in our model; however, since the amplitude of rhythmic clock gene expression in the hippocampus is modest, it is possible that even small changes in corticosterone profile or sampling times would prevent detection of differential gene expression in the current experimental design. Interestingly, in relation to phase-shifting properties of corticosterone, our data show that a 6 h delay in the corticosterone profile is accompanied by a similar shift in hippocampal Nr1d1, thus confirming that corticosterone independently governs hippocampal clock gene expression. In conclusion, circadian oscillations within physiological levels in serum corticosterone are sufficient to drive rhythmic expression of key constituents of the molecular clock in the hippocampus, suggesting that rhythmic glucocorticoids constitute a physiological link between the central clock of the SCN and the circadian oscillator of the hippocampus.

The authors wish to thank Rikke Lundorf and Tine Mellergaard (University of Copenhagen) for expert technical assistance. We also wish to express our gratitude to Dr. René Lemcke (University of Copenhagen) for expert guidance on the use of statistical software.

This study was approved by The Danish Council for Animal Experiments (authorization number 2017-15-0201-01190) and by the Faculty of Health and Medical Sciences at the University of Copenhagen (P21-190).

The authors report no conflict of interest.

This study was supported by the Lundbeck Foundation (grant number R344-2020–261 to MFR), Independent Research Fund Denmark (grant number 1030-00045B to MFR), the Novo Nordisk Foundation (grant number NNF21OC0070214 to MFR), Læge Sofus Carl Emil Friis og hustru Olga Doris Friis’ Legat (to MFR), and Dagmar Marshalls Fond (to MFR).

MFR conceived the study. TB and MFR designed experiments. TB, ASBV, and MFR performed experiments. TB, ASBV, and MFR analyzed data. TB and MFR wrote the manuscript. ASBV revised the manuscript. All authors approved the final manuscript.

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