BioMed Research International

BioMed Research International / 2017 / Article

Research Article | Open Access

Volume 2017 |Article ID 8459385 |

Jennifer Caroline Sousa, Keila Karoline Magalhães-Marques, Sanseray da Silveira Cruz-Machado, Maria Nathalia Moraes, Ana Maria de Lauro Castrucci, "Dexamethasone Modulates Nonvisual Opsins, Glucocorticoid Receptor, and Clock Genes in Danio rerio ZEM-2S Cells", BioMed Research International, vol. 2017, Article ID 8459385, 14 pages, 2017.

Dexamethasone Modulates Nonvisual Opsins, Glucocorticoid Receptor, and Clock Genes in Danio rerio ZEM-2S Cells

Academic Editor: Giuseppe Piccione
Received10 Oct 2016
Revised19 Mar 2017
Accepted22 Mar 2017
Published14 May 2017


Here we report, for the first time, the differential cellular distribution of two melanopsins (Opn4m1 and Opn4m2) and the effects of GR agonist, dexamethasone, on the expression of these opsins and clock genes, in the photosensitive D. rerio ZEM-2S embryonic cells. Immunopositive labeling for Opn4m1 was detected in the cell membrane whereas Opn4m2 labeling shows nuclear localization, which did not change in response to light. opn4m1, opn4m2, gr, per1b, and cry1b presented an oscillatory profile of expression in LD condition. In both DD and LD condition, dexamethasone (DEX) treatment shifted the peak expression of per1b and cry1b transcripts to ZT16, which corresponds to the highest opn4m1 expression. Interestingly, DEX promoted an increase of per1b expression when applied in LD condition but a decrease when the cells were kept under DD condition. Although DEX effects are divergent with different light conditions, the response resulted in clock synchronization in all cases. Taken together, these data demonstrate that D. rerio ZEM-2S cells possess a photosensitive system due to melanopsin expression which results in an oscillatory profile of clock genes in response to LD cycle. Moreover, we provide evidence that glucocorticoid acts as a circadian regulator of D. rerio peripheral clocks.

1. Introduction

Time is a fundamental variable for the manifestation of biological phenomena, and as such it has been systematically investigated since the pioneer chronobiology studies by Halberg [1]. The generation and maintenance of biological rhythms are explained based on a molecular machinery composed of positive and negative feedback loops of transcription and translation of core genes [29]. Although major work has been done with rodents, nonmammalian vertebrates have been explored by our group to favor an evolutionary perspective of the fundamentals of the biological clocks [1017].

Danio rerio, the popular zebrafish, has been employed as an alternative model to study clock molecular machinery, due to the similarities with the mammalian clock regarding the genes and their feedback loops [18, 19]. On the other hand, unlike mammals, the existence of photosensitive biological clocks in many peripheral organs [11, 20] allows the investigation of processes subjacent to light transduction in peripheral clocks.

The photopigments responsible for converting light into temporal information are assembled in a family of nonimage forming opsins named melanopsins, which were firstly discovered in Xenopus laevis melanophores [21], with subsequent description of orthologues in the retina of all vertebrate classes [2225], and extraocular tissues of some species [2628]. Data on genomic sequence similarity, chromosomal localization, and phylogeny revealed that melanopsin genes have evolved into two separate groups: Opn4m, orthologue of mammalian melanopsin, and Opn4x, orthologue of the melanopsin initially cloned from X. laevis [24, 25]. Both melanopsins are expressed in fish, amphibians, and birds, while mammals have apparently lost the ancestral Opn4x locus during chromosomal reorganization [25]. Surprisingly, in adult Danio rerio retina, five melanopsin genes, opn4m1, opn4m2, opn4m3, opn4x1, and opn4x2 [29], are found, although opn4m1 and opn4m2 are the mostly abundant melanopsins expressed in zebrafish embryonic cells (ZEM-2S) [14]. The high number of melanopsin genes is most probably due to teleost-specific whole genome duplication event which occurred in an ancient fish 350 million years ago [30].

Mammalian melanopsin is expressed by a small subset of retinal ganglion cells and translates light stimulus into neural information, which reaches the suprachiasmatic nucleus (SCN) through the retinohypothalamic tract [31, 32]. By contrast, the wide melanopsin expression in zebrafish peripheral tissues allows this fish to be directly entrained by light/dark cycles even in cultured cells [11, 14, 20, 3335].

The molecular mechanisms of zebrafish circadian rhythm are similar to the well-known mammalian system and rely on core set of genes composed of clock, bmal, per, and cry. Clock:Bmal dimer induces rhythmic expression of per and cry, while Per and Cry proteins repress their own expression by interacting with Clock:Bmal via the processes of blocking and displacement repressions [36]. In addition, there exists a stabilizing loop, which stimulates or represses bmal expression through the clock-controlled genes, Ror and Rev-erb, respectively. The completion of this feedback system takes about 24 h, thus generating a pattern of circadian rhythm [19].

However, some differences as the multiple clock genes and the light sensitivity make zebrafish a peculiar vertebrate model. Four period (per1a, per1b, per2, and per3), 6 cryptochrome (cry1a, cry1b, cry2a, cry2b, cry3, and cry4), 3 clock (clock1a, clock1b, and clock2), and 3 bmal (bmal1a, bmal1b, and bmal2) genes have been cloned from D. rerio [18, 19, 37, 38]. These multiple copies probably perform specialized functions in the clock regulation: all Cry proteins, except Cry3 and Cry4, and Per proteins inhibit Clock:Bmal [37]. The zebrafish light sensitivity is related to light induction of per2 and cry1a genes via D box-binding factor TEF (Thyrotroph Embryonic Factor) [3941], a signaling pathway which has recently been receiving the researchers’ attention [14, 19].

The synchronization between central and peripheral clocks raised the hypothesis that humoral factors are responsible for their communication. Hormones such as glucocorticoids (GCs) became strong candidates as regulatory agents of clock genes, as they are rhythmically produced and released in all vertebrates [4244]. One-hour pulse of dexamethasone (DEX), a synthetic glucocorticoid analogue, induces the circadian expression of Per1 in rat fibroblasts [45]. In fact, mammalian Per1 gene possesses the glucocorticoid responsive element (GRE), indicating that GC-receptor complex may directly modulate Per1 expression [46]. Mouse primary culture of mesenchymal cells responds with oscillatory transcription of Per1, Per2, Per3, Cry1, Cry2, Npas2 (Clock paralogue), and Bmal1 and of the clock-controlled genes, Rev-erb α/β and Dbp to GCs [47]. In rat hepatocyte primary culture, GCs also inhibit Rev-erbαtranscription [48].

Although the literature strongly indicates regulation of mammalian biological clocks by GCs, the mechanisms and the relevance of this modulation in nonmammalian vertebrates are yet to be established. In zebrafish corticotrope-deficient larvae, it has been demonstrated that dexamethasone was able to rescue high-amplitude circadian cell cycle rhythms [49].

Here, we investigated the putative role of GCs as synchronizing agents of Danio rerio embryonic cells ZEM-2S and the evoked regulation of melanopsins, GC receptor, per1b, and cry1b expression in this photosensitive cell line.

2. Material and Methods

2.1. ZEM-2S Cell Culture

Danio rerio embryonic cells (ZEM-2S) (gently donated by Professor Mark Rollag, Uniformed Services University of the Health Sciences, USA, and originally purchased from ATCC, CRL-2147, Manassas, VA, USA) were kept in 50% Leibowitz L-15, 35% Dulbecco’s Modified Eagle Medium (DMEM), 15% Ham’s F12 medium (Athena, Campinas, Brazil), and 15 mM 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Life Technologies, Carlsbad, CA, USA), supplemented with 10% fetal calf serum (Vitrocell, Campinas, SP, Brazil), 1% antibiotic/antimycotic solution (10,000 U penicillin/10,000 U streptomycin/25 µg amphotericin B, Life Technologies, Carlsbad, CA, USA), and all-trans-retinal at final concentration of 10−7 M (Sigma, St. Louis, MO, USA), pH 7.2, at 28°C.

For the experimental protocols the serum concentration was reduced to 2%. Cells were handled in the dark under a red safelight (7 W Konex bulb and Safelight Filter GBX-2, Kodak, Rochester, NY, USA).

2.2. Immunocytochemistry of Opn4m1 and Opn4m2

The 15 N-terminal sequences with an appended C-terminal cysteine of D. rerio Opn4m1 (MSGAAHSVRKGISC) and Opn4m2 (MSHHSSWRGHHCAPGC) were conjugated to keyhole limpet hemocyanin and inoculated into rabbits to obtain the primary antisera (Covance Labs, Denver, PA, USA). The primary antisera were previously tested in a panel of dilutions ranging from 1 : 100 to 1 : 1000, and good staining was observed only at 1 : 100 and 1 : 250 for both antibodies. The antisera and the Cy3-labeled secondary anti-rabbit antibody (Jackson ImmunoResearch, West Grove, PA, USA, 1 : 500) were diluted in incubation buffer (1% bovine serum albumin, 0.25% carrageenan lambda, and 0.3% Triton X-100, all from Sigma-Aldrich, St. Louis, MO, USA).

The expressions of Opn4m1 and Opn4m2 were evaluated in ZEM-2S cells kept in DD condition for 3 days and subject to 1 hour of white light pulse (650 lux) at the beginning of the 4th day. Control cells were kept in DD for the duration of the experiment. Twenty-four hours after the light pulse, cells were fixed with 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA) in phosphate buffered saline (PBS) at 4°C for 30 min. The blockade of nonspecific sites was made with 6% normal goat serum plus 22.52 mg/mL glycine (both from Sigma-Aldrich, St. Louis, MO, USA) in PBS at 4°C for 1 h, followed by incubation with the primary antibodies (anti-Opn4m1, 1 : 250, and anti-Opn4m2, 1 : 100) overnight at 4°C. The cells were then incubated with the secondary antibody for 1 h at room temperature, mounted with DAPI-Vectashield hard aqueous medium (Vector Laboratories, Burlingame, CA, USA) and coverslipped. Photographs were taken in an inverted fluorescence microscope Axiovert 40CFL (Zeiss, Oberkochen, Germany).

2.3. Image Flow Cytometry

To determine the expression of Opn4m1 and Opn4m2 proteins at light and dark phases, we quantified the fluorescence of cells using imaging flow cytometry (FlowSight, Amnis, EMD-Millipore, Seattle, WA, USA). ZEM-2S cells (2.106) were kept in LD cycle for 3 days and on the 4th day the cells were fixed with 4% paraformaldehyde at ZT4 and ZT16. The material was then subject to similar procedure as described in the immunocytochemistry section, incubated in anti-Opn4m1 or anti-Opn4m2 polyclonal antibodies at 1 : 250 and 1 : 100 dilutions, respectively, followed by incubation with a sheep FITC-conjugated anti-rabbit antibody at 1 : 200 dilution (Sigma-Aldrich, St. Louis, MO, USA). Negative controls were incubated with FITC-conjugated anti-rabbit antibody, in the absence of the primary antibody. The analysis of 10,000 cell counts per sample was performed by IDEAS software (Amnis, EMD-Millipore, Seattle, WA, USA) based on scatter plot of bright field area versus aspect ratio and expressed as normalized frequency based on FITC fluorescence intensity.

2.4. Temporal Expression of Melanopsin and Clock Genes

ZEM-2S cells (106 cells/25 cm2 flask) were seeded in 2% serum-supplemented medium and subject to DD or 12 : 12 LD cycle (lights-on: 9:00 h/lights-off: 21:00 h, 680 lux or 99.28 mW/cm2, 2.5 × 10−14 photons s−1 cm−2, full spectrum of white light Ecolume, 8 W cool white light bulb, model YZ8W/8 W cool white fluorescent tube, T5-8 W, and SCT) for at least 5 days, at 28°C. On the following day, total RNA was extracted every 4 h over a day. Each protocol was repeated at least twice, with two or three flasks per time point to obtain to 6.

2.5. Dexamethasone Assays
2.5.1. One DEX Pulse

ZEM-2S cells were kept in LD cycles and in the beginning of the dark phase of the 5th day the culture medium was replaced with medium containing  M DEX. Twelve hours later the culture medium was replaced by fresh medium, and on the 6th day total RNA was extracted every 4 h during 24 h.

2.5.2. Three DEX Pulses

ZEM-2S cells were kept in LD cycles and treated with three 12-hour pulses of  M DEX during the dark phase of the 3rd, 4th, and 5th days. After each 12-hour pulse the DEX containing medium was replaced with fresh medium. Twelve hours after the third DEX treatment, the culture medium was replaced by fresh medium, and on the 6th day total RNA was extracted every 4 h during 24 h.

2.5.3. Five DEX Pulses

ZEM-2S cells were kept in DD conditions and were treated with 2-hour pulses of 10−7 M DEX for 5 days starting at ZT 0. After each 2-hour pulse the DEX containing medium was replaced with fresh medium. On the 6th day total RNA was extracted every 4 h during 24 h.

The medium of the control cells was changed simultaneously with DEX-treated cells, totaling up to 2, 6, or 10 medium changes, respectively. DEX concentration and application time were based on Gilchrest and colleagues [48], who reported that plasma cortisol of rainbow trout (Oncorhynchus mykiss) peaks in the dark phase.

2.6. Total RNA Extraction, Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR), and Quantitative PCR (qPCR)

Total RNA was extracted with TRIzol Reagent (Ambion, Foster City, CA, USA) according to the manufacturer’s instructions. The RNA pellet was resuspended in DEPC H2O (diethylpyrocarbonate, Ambion, Foster City, CA, USA) and treated with DNAse according to the manufacturer’s protocol (turbo DNA-Free™, Ambion, Foster City, CA, USA). RNA concentration (OD260) was determined in a spectrophotometer (Nanodrop, Wilmington, DE, USA), and 2 µg was submitted to RT-PCR (SuperScript III, Invitrogen, Carlsbad, CA, USA) using random primers. The reaction protocol was as follows: 5 min at 65°C and 1 min at 4°C; after the enzyme addition, 5 min at 25°C and 50 min at 50°C; and 15 min at 70°C in a thermocycler (Eppendorf, Hauppauge, NY, USA).

Using TaqMan approach to simultaneously analyze the expression of per1b, cry1b, and ribosomal 18S RNA or opn4m1, gr, and 18S RNA we prepared a mix of primers (300 ηM for genes and 50 ηM for 18S RNA, Table 1), fluorescent probes (200 ηM for genes and 50 ηM for 18S RNA, Table 1), and Supermix 2X (Bio-Rad Laboratories, Hercules, CA, USA, or Life Technologies, Carlsbad, CA, USA), supplemented to final concentrations of 400 µM dNTPS, 6 mM MgCl2, and 0,1 U/µL Platinum Taq DNA polymerase (Life Technologies, SP, Brazil). Each experimental cDNA was run in triplicate in 96-well plates.

(access number)
Primers and probesFinal concentration






Reverse: 5′-TCTCCCCCGGGCCAC-3′300 nM

FAM = carboxyfluorescein; TexRd = Texas Red fluorophore; Cy5 = Cyanine 5 fluorophore; Hex = 6-carboxy-2,4,4,4,7,7-hexachlorofluorescein succinimidyl ester fluorophore; BHQ_1 = black hole quencher 1; BHQ_2 = black hole quencher 2.
Gene nomenclature according to for Homo sapiens; for Mus musculus; for Danio rerio.

For opn4m2 and 18S RNA, the qPCR reactions contained iQ™ SYBR® Green Supermix 2X (Bio-Rad Laboratories, Hercules, CA, USA) or SYBR® GreenER™ qPCR SuperMix for iCycler® 2X (Life Technologies, Carlsbad, CA, USA) and specific primers, in final concentrations of 300 ηM for opn4m2 and 50 ηM for 18S RNA, respectively, in independent solutions. Primers and probes for Danio rerio were designed based on sequences obtained from GenBank ( and synthesized by IDT (Coralville, IA, USA). 18S RNA was used to normalize the values of specific genes, since it did not vary among time points under the various experimental conditions and has a stable expression in many tissues of zebrafish [50].

Primers’ efficiency was determined using serial dilutions (1, 1 : 2, 1 : 4, 1 : 8, and 1 : 16) of a single cDNA sample. The triplicate mean CT of each gene was plotted in the -axis and the log of cDNA dilutions in the -axis. The efficiency for each primer pair was calculated according the equation: , in which corresponds to the inclination angle of the linear regression curve. Values between 90% and 110% were considered as indicators of appropriate efficiency.

All assays were performed in iCycler or iQ5 (Bio-Rad Laboratories, Hercules, CA, USA) thermocycler, with the following protocols: multiplex, 1 cycle of 7 min at 95°C followed by 45 cycles of 30 sec at 95°C and 30 sec at 60°C; SYBR Green, 2 min at 50°C, 8:30 min at 95°C, 45 cycles of 15 sec at 95°C, 1 min at 60°C, 1 min at 95°C, 1 min at 55°C, and 80 cycles of 10 sec at 55°C with a gradual increase of 0.5°C. Negative controls without templates were routinely included.

2.7. Statistical Analysis

The method was used to analyze relative changes in gene expression by qPCR. The CT was obtained comparing the number of cycles in control and experimental wells and among the various time points, by passing a threshold line through the geometric portions of the amplification curves. The was calculated as the difference between the CT for 18S RNA and that for the gene of interest; the smallest value obtained in the control group or in a time point was then subtracted from each , originating the . This value was then used as the negative exponential of base 2. To assess temporal variation among different time points (ZTs), means were compared by one way analysis of variance (ANOVA) followed by Tukey test; to compare control and experimental groups along time, two way ANOVA followed by Bonferroni test was used. All data were analyzed and graphed by GraphPad Prism 5 software, and the differences were considered statistically significant when .

3. Results and Discussion

3.1. Melanopsins and Glucocorticoid Receptor

The striking diversity of extra genes expressed in D. rerio explained by genome duplication event in the teleost lineage may reflect either redundant or specialized functions performed by the multiple copies [19]. However, the fact that the five melanopsin genes are differentially expressed in many neuronal cell types in the adult fish retina leads toward the presence of a finely tuned/sophisticated melanopsin system [29, 50].

By contrast, not all opn4 mRNAs were detected in earlier stages of development, as demonstrated in D. rerio ZEM-2S cells, where opn4m1 and opn4m2 genes are more expressed as compared to opn4m3, opn4x1, and opn4x2 genes, whose expression was negligible [14]. Thereby, in this study only opn4m1 and opn4m2 genes were analyzed besides the glucocorticoid receptor (gr) gene which was also found to be expressed in this cell line.

Here we demonstrate for the first time the presence and localization of the proteins Opn4m1 and Opn4m2 in D. rerio ZEM-2S cells using immunocytochemistry and imaging flow cytometry. Positive labeling for Opn4m1 (Figures 1(b) and 1(d)) was remarkably detected in the cell membrane at 1 : 250 dilution in cells kept in DD condition; no immunostaining was observed after incubation with rabbit preserum instead of the anti-Opn4m1 antiserum (Figure 1(a)). Interestingly Opn4m2 labeling is nuclear and much less evident in the membrane than Opn4m1 (Figures 2(b) and 2(d)). Control preparation incubated with preserum showed no immunolabeling (Figure 2(a)). Although it is commonly accepted that opsins are typically located in the cell membrane, similar results were also reported in Xenopus laevis melanophores [16]. These cells express two melanopsins, Opn4x and Opn4m, which are also differentially distributed in the cell: immunoreactivity for Opn4x was seen in the cytoplasm and cell membrane whereas Opn4m is strongly expressed in the nucleus [16]. The precise explanation for this distinct distribution is elusive and further investigation is needed to solve this question. However, a recent study using mammalian cells brought a possible explanation [51]. Mammals express only one melanopsin [25], and in melanocytes this opsin is located in the nucleus [51]. Interestingly after a 15 min white light pulse this opsin translocates to the cell membrane. In contrast, in ZEM-2S cells, light pulse was not able to induce the translocation of Opn4m2 to membrane (see Supplemental Figure in Supplementary Material available online at The differential response between nonmammalian and mammalian melanopsins may be due to the fact that mammals express only one melanopsin gene/protein, while D. rerio and X. laevis express multiple melanopsins. In D. rerio Opn4m1 found in the cell membrane likely works as a photosensor, while the nuclear melanopsin (Opn4m2) could function as a transcription factor. In addition, analyses of flow cytometry imaging revealed that the majority of the cells were positive for Opn4m1 (ZT4 91.9%   7.5% versus ZT16 99.07%   0.12%) and Opn4m2 (ZT4 99.75%   0.1% versus ZT16 99.43%   0.20%). However no difference in fluorescence intensity between ZTs was found (Figure 3). Taken together, these data suggest that although D. rerio expresses more than one melanopsin protein and there was no variation between light and dark phases, only Opn4m1, located in the membrane, may be responsible for transducing light stimulus.

LD cycle is a well-known zeitgeber that synchronize the clock components both in vitro and in vivo, allowing the rhythmic expression of several clock-controlled molecules. In this context, we analyzed the temporal profile of melanopsins and gr gene expression in LD cycles. ZEM-2S cells kept in LD cycles showed similar temporal variation of opn4m1 and opn4m2 genes which peaked in the dark phase at ZT16 (Figures 4(a) and 4(b)). Although opn4m1 transcript peaked at ZT16 (Figure 4(a)), no difference of protein expression was seen between ZTs (Figure 3), which could be happening in other ZTs. As to gr, its expression slightly decreased over the course of the 24 h; that is, higher gr expression was seen at ZT0 in comparison to ZT20 (Figure 4(c)).

The oscillatory profiles of both melanopsins in LD cycle (Figures 5(a) and 5(b)) were altered after 2 medium changes (control cells), the peak shifting to the light phase. Nevertheless, one DEX pulse was able to shift opn4m1 peak of expression back to the dark phase (Figure 5(a)), but not of opn4m2 (Figure 5(b)). Surprisingly, 6 medium changes (control cells for 3 hormone pulses) abolished the temporal oscillation of opn4m1 seen in LD before medium changes, and 3 pulses of DEX restored its temporal variation, defining a robust circadian rhythm, which peaked again during the dark phase (Figure 6(a)). For opn4m2, similar medium changes did not affect the oscillation seen in LD without manipulation (peak in the dark), and 3 pulses of DEX remarkably increased mRNA levels keeping the peak in the beginning of the dark phase (Figure 6(b)). These results demonstrate that glucocorticoids restored opn4m1 temporal profile in LD, which had been affected by medium changes, indicating that this opsin is more susceptible to dexamethasone treatment than opn4m2. Despite the fact that the glucocorticoid responsive element (GRE) has yet to be demonstrated in the melanopsin genes, its presence in opn4m1 but not in opn4m2 promoter might explain the differential response of these melanopsins to glucocorticoids.

Therefore, it is most likely that a functional circadian photopigment is present in these embryonic cells associated with the adjustment of the clock molecular machinery to light conditions. It was once thought that cry genes could exert a role in photoentrainment of vertebrate biological clocks because they mediate a variety of light responses, and their evolutionary history is related to photolyases [52]. In particular, cry1b and cry3 were indicated as potential candidates for this function in D. rerio Z3 cells [34]. More recently, however, we have demonstrated through pharmacological intervention in the rhabdomeric signaling pathways that melanopsins are the strongest candidate to exert the photopigment role in ZEM-2S cells [14]. Altogether, these results and the literature data strengthen the possibility that this opsin is the functional photopigment in D. rerio ZEM-2S cells.

The main mechanism of Gr regulation is performed by homologous downregulation, which occurs by decreasing the rate of gr transcription [53]. Our results confirm that dexamethasone affects the expression of its own receptor along 24 hours. One DEX pulse imposed a temporal variation with a peak of gr mRNA in the light phase (Figure 7(a)). Six medium changes or 3 pulses of DEX drastically increased the level of gr transcripts, respectively, around 50- and 80-fold, the values seen after 2 medium changes, and shifted the peak to the light-dark transition (Figure 7(b)).

In mammals, it is known that basal levels of glucocorticoid hormones exhibit marked circadian variation produced by secretory episodes with constant frequency and variable amplitude, which would exert an impact on the pattern of target gene transcription via GREs in target gene promoters [54]. Furthermore, nongenomic pathways have been described to be triggered by membrane-associated GR and second messengers [5560].

Indeed, since the last decade this hormone class has been related to circadian regulation of peripheral clocks [45, 47, 48, 6163]. The influence of DEX pulses on opn4m1 and opn4m2 expression shows that there may be a direct or indirect effect involving a signaling pathway evoked by Gr-glucocorticoid binding.

3.2. Clock Genes

The presence of multiple clock genes expressed in embryos and larvae of D. rerio and the ability to respond to light support the embryonic cell line ZEM-2S as a valuable model to study clock regulation. Although embryonic clocks are differentially regulated from those in the adult organism [64], photic signals and, consequently, an active phototransduction system, during blastula to early segmentation stages, are indispensable to maturation of a functional circadian clock [40].

Despite the relevance of peripheral clocks for understanding the organization of circadian system of an organism, the regulation of peripheral timing remains to be elucidated [65]. Hormones, whose temporal production and secretion are certainly defined by the master biological clock, have been particularly investigated as synchronizing agents of mammalian peripheral clocks [42, 66]. For nonmammalian vertebrates, the hormonal action on peripheral clocks has been less studied and its investigation could uncover evolutionary features of this regulation.

Circadian expression of clock genes in the presence of glucocorticoids has been evaluated in mammalian models of peripheral clocks. Balsalobre and coworkers [61] showed that 1-hour DEX-treatment was able to increase Per1 mRNA levels and promote a rhythmic expression of Per1, Per2, Per3, and Cry1 and clock-controlled genes Rev-erbαand Dbp in rat-1 fibroblasts. Dexamethasone downregulated Rev-erbαtranscripts in rat primary hepatocytes [48], and in cultured mouse liver cells, chronic prednisolone treatment induced Per1 mRNA expression and attenuated oscillatory profiles of Per2, Rev-erbα, and Bmal1 [62]. All PER and BMAL1 genes were also affected by glucocorticoids in human peripheral blood mononuclear cells [63].

Having this in mind we investigated the temporal expression profile of clock genes per1b and cry1b in the ZEM-2S cells in response to LD cycles and DEX treatment. Under LD cycles, mRNA levels of per1b presented a pronounced circadian oscillation with a remarkable 40-fold difference between light and dark phase transcripts (Figure 8(a)), peaking at ZT0 and gradually decreasing till the dark phase. For cry1b, in turn, a discrete temporal variation was observed with higher points of expression at the transitions of light-dark and dark-light phases (Figure 8(b)). Thus, comparing to the expression of opn4m1 and opn4m2 (Figures 4(a) and 4(b)), we found that the expression of per1b (Figure 8(a)) is in antiphase with melanopsin expression while no association was found for cry1b expression (Figure 9(b)).

Understanding the peripheral clock regulation in vivo is a complicated task, since several hormones may act to set the clock machinery. Interesting in nonmammalian cells that are able to directly detect light, hormones seem to play a minor role in the synchronization of clock components [15, 17]. Our results demonstrate a remarkable effect of dexamethasone pulses on clock genes. At ZT16 (dark phase) the expression of per1b (Figure 8(c)) and cry1b (Figure 8(d)) increased 30- and 50-fold, respectively, after 3 DEX pulses. This increase of per1b and cry1b in the dark phase coincides with a high expression of opn4m1 also after 3 DEX pulses.

The rhythmic profile of clock genes is clearly elicited by LD cycles since, in the absence of light, per1b exhibits a very slight variation, with a low amplitude peak during the subjective night (Figure 9(a)), while cry1b is constitutively expressed (Figure 9(b)). In a previous study, we have shown that medium change in DD induces a similar profile to LD, but with much lower amplitude [11]. A rhythm of lower amplitude has been shown for clock genes in zebrafish embryonic cell lines as PAC-2 (clock, [20]) and in Z3 cells (clock, per1, per3, bmal1, and bmal2, [39]), as well as in cultured zebrafish heart and kidney (clock [20]) maintained under constant conditions. The direct responsiveness to light stimulus is certainly the most remarkable characteristic of zebrafish cells. Similarly to per1b expression profile in ZEM-2S cells, per1 and per3 are rhythmically expressed in Z3 cells under LD cycles [39]. Phylogenetic studies revealed that the four per genes, per1a, per1b, per2 and per3, of D. rerio are, apparently, differentially regulated, with distinct spatial and temporal patterns [38]. In particular, per2 is light-inducible and does not oscillate in constant conditions [14, 18, 19].

We also investigated the ability of DEX to synchronize clock gene expression in the absence of light. Although per1b expression in cells subject to medium changes in DD seems to oscillate with higher values at ZTs 0, 4, and 8, no statistical differences were found (Figure 9(a)). DEX treatment in DD promoted a temporal oscillation of per1b transcription with high expression at ZT16 in comparison to ZTs 0, 4, 8, and 12. In addition, DEX induced a reduction of per1b expression at ZTs 0, 4, and 8, as compared to the untreated group. DEX treatment in DD (Figure 9(a)) did mimic LD cycle effects (Figure 8(a)) on per1b expression, inducing an oscillation, but with different profile. That is, in the first condition it peaked at ZT16 whereas in LD it was higher at ZT0. If we compare the effects of DEX in DD and in LD, new information arises from the results: in DD DEX decreased (Figure 9(a)) while in LD it increased (Figure 8(c)) per1b expression. On the other hand, cells kept in DD subject to medium changes showed a temporal variation of cry1b, with higher expression at ZTs 4 and 8 in comparison to ZT0. After DEX treatment we observed a shift in the peak expression of cry1b (Figure 9(b)) from ZTs 4 and 8 (control cells, medium changes) to ZTs 8 and 12. Unlike per1b, cry1b transcription was increased by DEX in both DD (Figure 9(b)) and LD (Figure 8(d)).

Glucocorticoids exhibit a strong daily rhythm in the plasma of many vertebrates from fish to mammals [6769]. This rhythmic glucocorticoid release is a signal to peripheral oscillators in mammalian liver, heart, lung, stomach, and kidney [46, 62]. However, the role of glucocorticoids in the regulation of clock gene expression in fish is still speculative. Sánchez-Bretaño and colleagues demonstrated, in a very elegant work in Carassius auratus, that, similarly to what we found in D. rerio cultured embryonic cells, DEX induces per genes in the liver in in vivo assays as well as in cultured hepatocytes [70]. In light responsive peripheral clocks, light has been demonstrated to prevail in regulating opsins and clock genes, as compared to many hormones, such as melatonin, α-MSH [13], and endothelin [15]. Despite the fact that D. rerio ZEM-2S cells are also photosensitive, glucocorticoids seem to be as relevant as light to regulate photoreception and the circadian system.

4. Conclusions

In summary, we report here, for the first time, the localization and daily variation of the melanopsin proteins, Opn4m1 and Opn4m2, in a nonmammalian photosensitive clock, the D. rerio ZEM-2S embryonic cells. It is interesting to mention that unlike mammals in which the single melanopsin migrates from the nucleus to the cell membrane in response to light, in ZEM-2S cells, the nuclear melanopsin does not relocate after light stimulus. We provide evidences of DEX stimulatory effect on all genes, except for per1b in ZEM-2S cells kept under DD. These results demonstrate the remarkable influence of glucocorticoids in all organizational levels of peripheral D. rerio circadian system, that is, photoperception, clock core, and clock-controlled genes such as gr.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Authors’ Contributions

Maria Nathalia Moraes and Ana Maria de Lauro Castrucci contributed equally to this work.


This work was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP, Grant 2012/50214-4) and Conselho Nacional de Desenvolvimento Tecnológico e Científico (CNPq, Grants 301293/2011-2 and 303070/2015-3) grants. Jennifer Caroline Sousa, Sanseray da Silveira Cruz-Machado, and Maria Nathalia Moraes are fellows of FAPESP.

Supplementary Materials

Danio rerio possess five melanopsin genes (Davis et al., 2011); among them, opn4m1 and opn4m2 were reported as the most expressed melanopsins in ZEM-2S cells (Ramos et al., 2014). The cellular compartments where Opn4m1 and Opn4m2 are expressed were identified here by immunocytochemistry. Surprisingly Opn4m2 was found in the nucleus when ZEM-2S cells were kept in DD conditions. So, we investigated the effects of white light pulse in promoting melanopsin translocation. Danio rerio ZEM-2S cells were kept for 3 days in DD condition and subject to 1 hour of white light pulse (650 lux) in the beginning of the 4th day. The incubation of ZEM-2S cells with anti-Opn4m2 at 1 : 100 dilution demonstrated that white light pulse did not altered Opn4m2 nuclear location. On the other hand, in mammalian melanoma cells, a short white light pulse (15 min) promotes melanopsin translocation from the nucleus to the cell membrane (de Assis et al., 2016). Davies, W. I. L. et al. (2011). Functional diversity of melanopsins and their global expression in the teleost retina. Cell. Mol. Life Sci. 68(24), 4115–4132. Ramos, B. C. R., Moraes, M. N. C. M., Poletini, M. O., Lima, L. H. R. G., and Castrucci, A. M. L. (2014) From blue light to clock genes in zebrafish ZEM-2S cells. PLoS ONE 9 (9), Article ID e106252. de Assis, L. V. M., Moraes, M. N., da Silveira Cruz-Machado, S., and Castrucci, A. M. L. (2016) The effect of white light on normal and malignant murine melanocytes: a link between opsins, clock genes, and melanogenesis. Biochim. Biophys. Acta 1863(6), 1119–1133.

  1. Supplementary Material


  1. F. Halberg, “Chronobiology.,” Annual Review of Physiology, vol. 31, pp. 675–725, 1969. View at: Publisher Site | Google Scholar
  2. M. H. Vitaterna, D. P. King, A.-M. Chang et al., “Mutagenesis and mapping of a mouse gene, clock, essential for circadian behavior,” Science, vol. 264, no. 5159, pp. 719–725, 1994. View at: Publisher Site | Google Scholar
  3. H. Tei, H. Okamura, Y. Shigeyoshi et al., “Circadian oscillation of a mammalian homologue of the Drosophila period gene,” Nature, vol. 389, no. 6650, pp. 512–516, 1997. View at: Publisher Site | Google Scholar
  4. T. Takumi, C. Matsubara, Y. Shigeyoshi et al., “A new mammalian period gene predominantly expressed in the suprachiasmatic nucleus,” Genes to Cells, vol. 3, no. 3, pp. 167–176, 1998. View at: Publisher Site | Google Scholar
  5. T. Takumi, K. Taguchi, S. Miyake et al., “A light-independent oscillatory gene mPer3 in mouse SCN and OVLT,” The EMBO Journal, vol. 17, no. 16, pp. 4753–4759, 1998. View at: Publisher Site | Google Scholar
  6. N. Gekakis, D. Staknis, H. B. Nguyen et al., “Role of the CLOCK protein in the mammalian circadian mechanism,” Science, vol. 280, no. 5369, pp. 1564–1569, 1998. View at: Publisher Site | Google Scholar
  7. G. T. J. Van Der Horst, M. Muijtjens, K. Kobayashi et al., “Mammalian Cry1 and Cry2 are essential for maintenance of circadian rhythms,” Nature, vol. 398, no. 6728, pp. 627–630, 1999. View at: Publisher Site | Google Scholar
  8. S. Yamazaki, R. Numano, M. Abe et al., “Resetting central and peripheral circadian oscillators in transgenic rats,” Science, vol. 288, no. 5466, pp. 682–685, 2000. View at: Publisher Site | Google Scholar
  9. U. Albrecht and G. Eichele, “The mammalian circadian clock,” Current Opinion in Genetics & Development, vol. 13, no. 3, pp. 271–277, 2003. View at: Google Scholar
  10. L. H. R. G. Lima, A. C. Scarparo, M. C. Isoldi, M. A. Visconti, and A. M. L. Castrucci, “Melanopsin in chicken melanocytes and retina,” Biological Rhythm Research, vol. 37, no. 5, pp. 393–404, 2006. View at: Publisher Site | Google Scholar
  11. F. P. Farhat, C. B. Martins, L. H. Ribeiro Graciani De Lima, M. C. Isoldi, and A. M. D. L. Castrucci, “Melanopsin and clock genes: regulation by light and endothelin in the zebrafish ZEM-2S cell line,” Chronobiology International, vol. 26, no. 6, pp. 1090–1119, 2009. View at: Publisher Site | Google Scholar
  12. L. H. R. G. De Lima, K. P. Dos Santos, and A. M. De Lauro Castrucci, “Clock genes, melanopsins, melatonin, and dopamine key enzymes and their modulation by light and glutamate in chicken embryonic retinal cells,” Chronobiology International, vol. 28, no. 2, pp. 89–100, 2011. View at: Publisher Site | Google Scholar
  13. M. N. D. C. M. Moraes, L. R. D. Santos, N. Mezzalira, M. O. Poletini, and A. M. D. L. Castrucci, “Regulation of melanopsins and per1 by α -MSH and melatonin in photosensitive xenopus laevis melanophores,” BioMed Research International, vol. 2014, Article ID 654710, pp. 1–10, 2014. View at: Publisher Site | Google Scholar
  14. B. C. R. Ramos, M. N. C. M. Moraes, M. O. Poletini, L. H. R. G. Lima, and A. M. L. Castrucci, “From blue light to clock genes in zebrafish ZEM-2S cells,” PLoS ONE, vol. 9, no. 9, Article ID e106252, 2014. View at: Publisher Site | Google Scholar
  15. M. N. D. C. M. Moraes, L. H. R. G. D. Lima, B. C. R. Ramos, M. D. O. Poletini, and A. M. D. L. Castrucci, “Endothelin modulates the circadian expression of non-visual opsins,” General and Comparative Endocrinology, vol. 205, pp. 279–286, 2014. View at: Publisher Site | Google Scholar
  16. M. N. Moraes, B. C. Ramos, M. O. Poletini, and A. M. L. Castrucci, “Melanopsins: localization and Phototransduction in Xenopus laevis Melanophores,” Photochemistry and Photobiology, vol. 91, no. 5, pp. 1133–1141, 2015. View at: Publisher Site | Google Scholar
  17. M. O. Poletini, B. C. Ramos, M. N. Moraes, and A. M. L. Castrucci, “Nonvisual opsins and the regulation of peripheral clocks by light and hormones,” Photochemistry and Photobiology, vol. 91, no. 5, pp. 1046–1055, 2015. View at: Publisher Site | Google Scholar
  18. G. M. Cahill, “Clock mechanisms in zebrafish,” Cell and Tissue Research, vol. 309, no. 1, pp. 27–34, 2002. View at: Publisher Site | Google Scholar
  19. G. Vatine, D. Vallone, Y. Gothilf, and N. S. Foulkes, “It's time to swim! Zebrafish and the circadian clock,” FEBS Letters, vol. 585, no. 10, pp. 1485–1494, 2011. View at: Publisher Site | Google Scholar
  20. D. Whitmore, N. S. Foulkes, and P. Sassone-Corsi, “Light acts directly on organs and cells in culture to set the vertebrate circadian clock,” Nature, vol. 404, no. 6773, pp. 87–91, 2000. View at: Publisher Site | Google Scholar
  21. I. Provencio, G. Jiang, W. J. De Grip, W. Pär Hayes, and M. D. Rollag, “Melanopsin: an opsin in melanophores, brain, and eye,” Proceedings of the National Academy of Sciences of the United States of America, vol. 95, no. 1, pp. 340–345, 1998. View at: Publisher Site | Google Scholar
  22. I. Provencio, I. R. Rodriguez, G. Jiang, W. P. Hayes, E. F. Moreira, and M. D. Rollag, “A novel human opsin in the inner retina,” Journal of Neuroscience, vol. 20, no. 2, pp. 600–605, 2000. View at: Google Scholar
  23. I. Provencio, M. D. Rollag, and A. M. L. Castrucci, “Photoreceptive net in the mammalian retina. This mesh of cells may explain how some blind mice can still tell day from nigh,” Nature, vol. 415, no. 6871, article 493, 2002. View at: Google Scholar
  24. J. Bellingham, D. Whitmore, A. R. Philp, D. J. Wells, and R. G. Foster, “Zebrafish melanopsin: isolation, tissue localisation and phylogenetic position,” Molecular Brain Research, vol. 107, no. 2, pp. 128–136, 2002. View at: Publisher Site | Google Scholar
  25. J. Bellingham, S. S. Chaurasia, Z. Melyan et al., “Evolution of melanopsin photoreceptors: discovery and characterization of a new melanopsin in nonmammalian vertebrates,” PLoS Biology, vol. 4, no. 8, article no. e254, pp. 1334–1343, 2006. View at: Publisher Site | Google Scholar
  26. M. J. Bailey and V. M. Cassone, “Melanopsin expression in the chick retina and pineal gland,” Molecular Brain Research, vol. 134, no. 2, pp. 345–348, 2005. View at: Publisher Site | Google Scholar
  27. E. Frigato, D. Vallone, C. Bertolucci, and N. S. Foulkes, “Isolation and characterization of melanopsin and pinopsin expression within photoreceptive sites of reptiles,” Naturwissenschaften, vol. 93, no. 8, pp. 379–385, 2006. View at: Publisher Site | Google Scholar
  28. N. Cavallari, E. Frigato, D. Vallone et al., “A blind circadian clock in cavefish reveals that opsins mediate peripheral clock photoreception,” PLoS Biology, vol. 9, no. 9, Article ID e1001142, 2011. View at: Publisher Site | Google Scholar
  29. W. I. L. Davies, L. Zheng, S. Hughes et al., “Functional diversity of melanopsins and their global expression in the teleost retina,” Cellular and Molecular Life Sciences, vol. 68, no. 24, pp. 4115–4132, 2011. View at: Publisher Site | Google Scholar
  30. D. Alsop and M. Vijayan, “The zebrafish stress axis: molecular fallout from the teleost-specific genome duplication event,” General and Comparative Endocrinology, vol. 161, no. 1, pp. 62–66, 2009. View at: Publisher Site | Google Scholar
  31. D. M. Berson, F. A. Dunn, and M. Takao, “Phototransduction by retinal ganglion cells that set the circadian clock,” Science, vol. 295, no. 5557, pp. 1070–1073, 2002. View at: Publisher Site | Google Scholar
  32. S. Hattar, H.-W. Liao, M. Takao, D. M. Berson, and K.-W. Yau, “Melanopsin-containing retinal ganglion cells: architecture, projections, and intrinsic photosensitivity,” Science, vol. 295, no. 5557, pp. 1065–1070, 2002. View at: Publisher Site | Google Scholar
  33. D. Whitmore, N. S. Foulkes, U. Strähle, and P. Sassone-Corsi, “Zebrafish Clock rhythmic expression reveals independent peripheral circadian oscillators,” Nature Neuroscience, vol. 1, no. 8, pp. 701–707, 1998. View at: Publisher Site | Google Scholar
  34. N. Cermakian, M. P. Pando, C. L. Thompson et al., “Light induction of a vertebrate clock gene involves signaling through blue-light receptors and MAP kinases,” Current Biology, vol. 12, no. 10, pp. 844–848, 2002. View at: Publisher Site | Google Scholar
  35. A.-J. F. Carr and D. Whitmore, “Imaging of single light-responsive clock cells reveals fluctuating free-running periods,” Nature Cell Biology, vol. 7, no. 3, pp. 319–321, 2005. View at: Publisher Site | Google Scholar
  36. R. Ye, C. P. Selby, Y. Y. Chiou, I. Ozkan-Dagliyan, S. Gaddameedhi, and A. Sancar, “Dual modes of CLOCK:BMAL1 inhibition mediated by Cryptochrome and Period proteins in the mammalian circadian clock,” Genes & Development, vol. 28, no. 18, pp. 1989–1998, 2014. View at: Google Scholar
  37. Y. Kobayashi, T. Ishikawa, J. Hirayama et al., “Molecular analysis of zebrafish photolyase/cryptochrome family: two types of cryptochromes present in zebrafish,” Genes to Cells, vol. 5, no. 9, pp. 725–738, 2000. View at: Publisher Site | Google Scholar
  38. H. Wang, “Comparative analysis of period genes in teleost fish genomes,” Journal of Molecular Evolution, vol. 67, no. 1, pp. 29–40, 2008. View at: Publisher Site | Google Scholar
  39. M. P. Pando, A. B. Pinchak, N. Cermakian, and P. Sassone-Corsi, “A cell-based system that recapitulates the dynamic light-dependent regulation of the vertebrate clock,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 18, pp. 10178–10183, 2001. View at: Publisher Site | Google Scholar
  40. L. Ziv and Y. Gothilf, “Circadian time-keeping during early stages of development,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 11, pp. 4146–4151, 2006. View at: Publisher Site | Google Scholar
  41. D. Gavriouchkina, S. Fischer, T. Ivacevic, J. Stolte, V. Benes, and M. P. S. Dekens, “Thyrotroph embryonic factor regulates light-induced transcription of repair genes in zebrafish embryonic cells,” PLoS ONE, vol. 5, no. 9, Article ID e12542, pp. 1–10, 2010. View at: Publisher Site | Google Scholar
  42. T. Dickmeis, B. D. Weger, and M. Weger, “The circadian clock and glucocorticoids—interactions across many time scales,” Molecular and Cellular Endocrinology, vol. 380, no. 1-2, pp. 2–15, 2013. View at: Publisher Site | Google Scholar
  43. K. L. Gamble, R. Berry, S. J. Frank, and M. E. Young, “Circadian clock control of endocrine factors,” Nature Reviews Endocrinology, vol. 10, no. 8, pp. 466–475, 2014. View at: Publisher Site | Google Scholar
  44. A. H. Tsang, J. L. Barclay, and H. Oster, “Interactions between endocrine and circadian systems,” Journal of Molecular Endocrinology, vol. 52, no. 1, pp. R1–R16, 2013. View at: Publisher Site | Google Scholar
  45. A. Balsalobre, L. Marcacci, and U. Schibler, “Multiple signaling pathways elicit circadian gene expression in cultured Rat-1 fibroblasts,” Current Biology, vol. 10, no. 20, pp. 1291–1294, 2000. View at: Publisher Site | Google Scholar
  46. T. Yamamoto, Y. Nakahata, M. Tanaka et al., “Acute physical stress elevates mouse Period1 mRNA expression in mouse peripheral tissues via a glucocorticoid-responsive element,” Journal of Biological Chemistry, vol. 280, no. 51, pp. 42036–42043, 2005. View at: Publisher Site | Google Scholar
  47. A. Y.-L. So, T. U. Bernal, M. L. Pillsbury, K. R. Yamamoto, and B. J. Feldman, “Glucocorticoid regulation of the circadian clock modulates glucose homeostasis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 41, pp. 17582–17587, 2009. View at: Publisher Site | Google Scholar
  48. I. P. Torra, V. Tsibulsky, F. Delaunay et al., “Circadian and glucocorticoid regulation of Rev-erbα expression in liver,” Endocrinology, vol. 141, no. 10, pp. 3799–3806, 2000. View at: Publisher Site | Google Scholar
  49. T. Dickmeis, K. Lahiri, G. Nica et al., “Glucocorticoids play a key role in circadian cell cycle rhythms,” PLoS Biology, vol. 5, no. 4, pp. 854–864, 2007. View at: Publisher Site | Google Scholar
  50. A. T. McCurley and G. V. Callard, “Characterization of housekeeping genes in zebrafish: male-female differences and effects of tissue type, developmental stage and chemical treatment,” BMC Molecular Biology, vol. 9, article 102, 2008. View at: Publisher Site | Google Scholar
  51. L. V. M. de Assis, M. N. Moraes, S. da Silveira Cruz-Machado, and A. M. L. Castrucci, “The effect of white light on normal and malignant murine melanocytes: a link between opsins, clock genes, and melanogenesis,” Biochimica et Biophysica Acta, vol. 1863, no. 6, pp. 1119–1133, 2016. View at: Publisher Site | Google Scholar
  52. A. R. Cashmore, J. A. Jarillo, Y.-J. Wu, and D. Liu, “Cryptochromes: blue light receptors for plants and animals,” Science, vol. 284, no. 5415, pp. 760–765, 1999. View at: Publisher Site | Google Scholar
  53. T. P. Mommsen, M. M. Vijayan, and T. W. Moon, “Cortisol in teleosts: dynamics, mechanisms of action, and metabolic regulation,” Reviews in Fish Biology and Fisheries, vol. 9, no. 3, pp. 211–268, 1999. View at: Publisher Site | Google Scholar
  54. T. Dickmeis and N. S. Foulkes, “Glucocorticoids and circadian clock control of cell proliferation: at the interface between three dynamic systems,” Molecular and Cellular Endocrinology, vol. 331, no. 1, pp. 11–22, 2011. View at: Publisher Site | Google Scholar
  55. J. W. Funder, “Glucocorticoid receptors,” Journal of Steroid Biochemistry and Molecular Biology, vol. 43, no. 5, pp. 389–394, 1992. View at: Publisher Site | Google Scholar
  56. S. L. Lightman, R. J. Windle, M. D. Julian et al., “Significance of pulsatility in the HPA axis,” in Proceedings of the Novartis Foundation Symposia, vol. 227, pp. 244–257, 2000. View at: Google Scholar
  57. S. L. Lightman, R. J. Windle, X.-M. Ma et al., “Hypothalamic-pituitary-adrenal function,” Archives of Physiology and Biochemistry, vol. 110, no. 1-2, pp. 90–93, 2002. View at: Publisher Site | Google Scholar
  58. S. L. Lightman, C. C. Wiles, H. C. Atkinson et al., “The significance of glucocorticoid pulsatility,” European Journal of Pharmacology, vol. 583, no. 2-3, pp. 255–262, 2008. View at: Publisher Site | Google Scholar
  59. T. Rhen and J. A. Cidlowski, “Antiinflammatory action of glucocorticoids—new mechanisms for old drugs,” New England Journal of Medicine, vol. 353, no. 16, pp. 1658–1723, 2005. View at: Publisher Site | Google Scholar
  60. A. McMaster, M. Jangani, P. Sommer et al., “Ultradian cortisol pulsatility encodes a distinct, biologically important signal,” PLoS ONE, vol. 6, no. 1, Article ID e15766, 2011. View at: Publisher Site | Google Scholar
  61. A. Balsalobre, S. A. Brown, L. Marcacci et al., “Resetting of circadian time in peripheral tissues by glucocorticoid signaling,” Science, vol. 289, no. 5488, pp. 2344–2347, 2000. View at: Publisher Site | Google Scholar
  62. S. Koyanagi, S. Okazawa, Y. Kuramoto et al., “Chronic treatment with prednisolone represses the circadian oscillation of clock gene expression in mouse peripheral tissues,” Molecular Endocrinology, vol. 20, no. 3, pp. 573–583, 2006. View at: Publisher Site | Google Scholar
  63. M. Cuesta, N. Cermakian, and D. B. Boivin, “Glucocorticoids entrain molecular clock components in human peripheral cells,” The FASEB Journal, vol. 29, no. 4, pp. 1360–1370, 2015. View at: Publisher Site | Google Scholar
  64. M. P. S. Dekens and D. Whitmore, “Autonomous onset of the circadian clock in the zebrafish embryo,” The EMBO Journal, vol. 27, no. 20, pp. 2757–2765, 2008. View at: Publisher Site | Google Scholar
  65. J. Husse, G. Eichele, and H. Oster, “Synchronization of the mammalian circadian timing system: Light can control peripheral clocks independently of the SCN clock: Alternate routes of entrainment optimize the alignment of the body's circadian clock network with external time,” BioEssays, vol. 37, no. 10, pp. 1119–1128, 2015. View at: Publisher Site | Google Scholar
  66. P. Pezük, J. A. Mohawk, L. A. Wang, and M. Menaker, “Glucocorticoids as entraining signals for peripheral circadian oscillators,” Endocrinology, vol. 153, no. 10, pp. 4775–4783, 2012. View at: Publisher Site | Google Scholar
  67. T. Ota, J.-M. Fustin, H. Yamada, M. Doi, and H. Okamura, “Circadian clock signals in the adrenal cortex,” Molecular and Cellular Endocrinology, vol. 349, no. 1, pp. 30–37, 2012. View at: Publisher Site | Google Scholar
  68. A. Kalsbeek and E. Fliers, “Daily regulation of hormone profiles,” in Handbook of Experimental Pharmacolology, vol. 217, pp. 185–226, 2013. View at: Google Scholar
  69. J. F. López-Olmeda, B. Blanco-Vives, I. M. Pujante, Y. S. Wunderink, J. M. Mancera, and F. J. Sánchez-Vázquez, “Daily rhythms in the hypothalamus-pituitary-interrenal axis and acute stress responses in a teleost flatfish, solea senegalensis,” Chronobiology International, vol. 30, no. 4, pp. 530–539, 2013. View at: Publisher Site | Google Scholar
  70. A. Sánchez-Bretaño, M. Callejo, M. Montero, Á. L. Alonso-Gómez, M. J. Delgado, and E. Isorna, “Performing a hepatic timing signal: glucocorticoids induce gper1a and gper1b expression and repress gclock1a and gbmal1a in the liver of goldfish,” Journal of Comparative Physiology B: Biochemical, Systemic, and Environmental Physiology, vol. 186, no. 1, pp. 73–82, 2016. View at: Publisher Site | Google Scholar

Copyright © 2017 Jennifer Caroline Sousa et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Related articles

No related content is available yet for this article.
 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder

Related articles

No related content is available yet for this article.

Article of the Year Award: Outstanding research contributions of 2021, as selected by our Chief Editors. Read the winning articles.