Biol Reprod
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

BOR - Papers in Press, published online ahead of print June 28, 2006.
Biol Reprod 2006, 10.1095/biolreprod.106.050732
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
75/4/624    most recent
biolreprod.106.050732v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Karman, B. N.
Right arrow Articles by Tischkau, S. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Karman, B. N.
Right arrow Articles by Tischkau, S. A.
Agricola
Right arrow Articles by Karman, B. N.
Right arrow Articles by Tischkau, S. A.
BIOLOGY OF REPRODUCTION 75, 624–632 (2006)
DOI: 10.1095/biolreprod.106.050732
© 2006 by the Society for the Study of Reproduction, Inc.

Research Article

Circadian Clock Gene Expression in the Ovary: Effects of Luteinizing Hormone1

Bethany N. Karman 3, and Shelley A. Tischkau 2 34 

Department of Veterinary Biosciences,3 Neuroscience Program,4 University of Illinois at Urbana-Champaign, Urbana, Illinois 61802

ABSTRACT

A molecular device that measures time on a daily, or circadian, scale is a nearly ubiquitous feature of eukaryotic organisms. A core group of clock genes, whose coordinated function is required for this timekeeping, is expressed both in the central clock and within numerous peripheral organs. We examined expression of clock genes in the rat ovary. Transcripts for core oscillator elements (Arntl,Clock,Per1,Per2, andCry1) were present in the ovary as indicated by quantitative real-time RT-PCR. Rhythmic expression patterns ofArntlandPer2transcripts and protein products were out of phase with respect to the central oscillator and in complete antiphase to each other. Expression ofArntlwas significantly elevated after the LH surge on the day of proestrus. Finally, hCG treatment induced cyclic expression of bothArntlandPer2gene products in hypophysectomized, immature rats primed with eCG. Collectively, these data suggest that the core underpinnings of the transcriptional/translational feedback loop that drives circadian rhythmicity is present in the rat ovary. Furthermore, the study identifies LH as a potential regulator of circadian clock gene rhythms in the ovary.

circadian rhythm, gene regulation, luteinizing hormone, mechanisms of hormone action, ovary

INTRODUCTION

Endogenous circadian rhythms measure time on a scale of ~24 h and orchestrate myriad physiological processes to allow organismic synchrony with the external environment. In mammals, entrainment to alternating periods of environmental light and darkness occurs at the level of the suprachiasmatic nucleus (SCN) of the basal hypothalamus primarily through glutamatergic neurotransmission via the retinohypothalamic tract [1–7]. Cellular circadian rhythms are generated by a highly conserved molecular mechanism comprised of a core group of clock genes and their protein products. These molecular components form interlocking transcriptional/translational autoregulatory feedback loops that oscillate with a near 24-h periodicity [8–11]. The transcription factor, aryl hydrocarbon receptor nuclear translocator-like (ARNTL; also frequently referred to as BMAL1 or MOP3), provides positive drive to the system by forming a heterodimer with CLOCK through protein-protein interactions between their period-arnt-single minded (PAS) domains. This ARNTL/CLOCK heterodimer drives transcription of cryptochrome (Cry1 and Cry2), Period (Pers 1, 2, and 3), and Timeless genes through binding to E-box elements in their promoter regions [8–11]. PER and CRY proteins undergo posttranslational modifications, form multimeric complexes and feed back to inhibit their own transcription [12]. In addition, PER2 acts as a positive regulator for Arntl [13, 14].

Circadian oscillations are present in virtually all tissues. Microarray analysis has revealed that 2%–10% of all genes exhibit circadian oscillations, but that rhythmicity of individual genes is tissue-specific, with less than 5% overlap between tissues [15]. Clock genes are rhythmically expressed in various peripheral tissues, including liver, muscle, kidney, and heart [16–24]. Regulation of molecular circadian rhythmicity in peripheral tissues remains enigmatic. The SCN likely provides an important source of the timing signal. Peak expression of Per2 and Arntl in liver, muscle, and kidney occurs in late subjective day and late subjective night, respectively, contrasting significantly with their corresponding peaks in the SCN. Furthermore, SCN ablation destroys rhythms of Per1, Per2, and Arntl in the liver [22, 25]. Despite recent evidence that peripheral organs may retain rhythmic features in vitro without the SCN [26], the overwhelming majority of data suggests that the SCN is required to synchronize cellular oscillations within a tissue.

The SCN plays an important role in regulating timing of the LH surge within the female reproductive axis. SCN-lesioned female rats lack the ovulatory LH surge and fail to ovulate [27]. Although it occurs only once every 4 or 5 days in the intact rat, the LH surge is precisely timed to occur on the afternoon of proestrus [28, 29]; daily LH surges are present in ovariectomized, estrogen-treated rats [28, 29]. Furthermore, external stimuli that alter the timing of the circadian activity rhythm likewise have the same effect on the timing of the LH surge [30]. This study examines events downstream of the LH surge to begin to determine whether the circadian–regulated LH surge may participate in regulation of circadian clock gene rhythms in the ovary.

Investigation of the role of clock genes in reproductive function remains in its infancy. Several studies in the mouse testis failed to detect rhythmic expression of Arntl, Per1, and Cry1 [31–34]; other studies revealed clear oscillations of testicular Per1, Per3, and Arntl [18, 35, 36]. Within the female reproductive system, Arntl and Per2 transcripts are rhythmically expressed in the rat oviduct [37]. However, clock gene expression in the ovary has not been investigated. This study begins to characterize expression and regulation of core clock genes in the ovary. We hypothesized that oscillations in core elements of the molecular clock are present in the ovary and proposed that the LH surge may regulate expression of one or more clock genes. Temporal expression patterns of Arntl and Per2 transcripts over a 24-h period and on 2 disparate days of the rat estrous cycle revealed daily oscillatory patterns of expression. Clock gene rhythms were absent in hypophysectomized, immature rats treated only with eCG. However, rhythmicity was induced in these animals upon treatment with LH (hCG). Arntl and Per2 expression patterns are consistent with the presence of functional molecular clockwork machinery in the ovary, and suggest that LH may play a role in regulating ovarian clock gene rhythms through actions on Arntl and Per2.

MATERIALS AND METHODS

Intact Adult Experiments

Eight-week-old female Sprague-Dawley rats (Harlan, Indianapolis, IN) were maintained in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, and all protocols were approved by the University of Illinois Animal Care and Use Committee. Food and water were provided ad libitum. Rats were housed under a strict 12L:12D lighting schedule. Under these lighting conditions, denoted as LD, zeitgeber time 0 (ZT 0) represents the time of lights-on in the colony. The corresponding time of lights-off is denoted as ZT 12. Animals were allowed to entrain to this lighting schedule for 2 wk before experimentation. Animals were killed by decapitation every 4 h over a 24-h period (n = 4 per group) starting at ZT 0. Ovaries were dissected from the animals and snap frozen in liquid nitrogen for immunoblot or mRNA analysis, or placed in 10% neutral buffered formalin for immunohistochemistry (IHC) and in situ hybridization. For analysis of mRNA and protein from SCN, hypothalamic brain slices (500 µM) containing the paired SCN were trimmed under a dissecting microscope to preserve only the SCN and underlying optic chiasm as described previously [38]. These reduced slices retain the electrophysiological and biochemical properties observed in full-size SCN slices [39]. Samples were frozen immediately in liquid nitrogen and stored at –80°C until processed for quantitative real-time RT-PCR (qPCR) or immunoblot analysis.

Estrous cycle stage was assessed from vaginal smears obtained every morning at ZT 4. Diestrus I was defined as vaginal smears containing abundant leukocytes mixed with cornified cells that appeared the day following a smear comprised entirely of cornified cells. Proestrus was defined as vaginal smears containing mostly small, nucleated epithelial cells, singly or in sheets, and the absence of leukocytes. Animals were monitored for at least two full estrous cycles before proceeding with the study, and those with irregular estrous cycles were omitted. After determination of cycle stage, animals were killed every 4 h over a 24-h period beginning at ZT 6 (2 h after staging) and ending at ZT 2 the following day (n = 3 per treatment group).

Hypophysectomized Immature Female Experiments

To examine the effects of gonadotropins on ovarian clock components, juvenile hypophysectomized animals were used. Female Sprague-Dawley rats were hypophysectomized at 20 days of age at Charles River Laboratories (Wilmington, MA). Animals arrived at 27 days of age, and were allowed to acclimate to a strict 12L:12D schedule for 48 h before treatment. Glucose (5%) was provided in the drinking water and along with food ad libitum. Animals were randomly assigned to hCG treatment and control groups (eCG followed by vehicle). On Day 30, both groups were administered 10 IU eCG in PBS (s.c.; Sigma, St. Louis, MO) to induce follicular development [40]. After 52 h, the treatment group was administered 10 IU hCG in PBS and controls received vehicle. Animals were killed by decapitation 0, 4, 8, 12, and 24 h after hCG treatment. Postmortem assessment was performed to confirm complete hypophysectomy (n = 3–5 per group).

Quantitative Real-time RT-PCR

After disruption and homogenization of the tissue, total ovarian or SCN RNA was extracted using TRIzol reagent according to the manufacturer's protocol (Invitrogen, Carlsbad, CA). Concentration, purity, and quality of the RNA were determined using the NanoDrop ND-1000 UV spectrophotometer at 260 nm and 280 nm and by gel electrophoresis. Total RNA (1 µg) was reverse transcribed using 200 ng random hexamers, 200 U SSII reverse transcriptase, 10 mM DTT, and 1x First-Strand Buffer (Invitrogen) at 43°C for 1 h in a volume of 20 µl (Mycycler; Bio-Rad, Hercules, CA). Negative controls included omission of reverse transcriptase and omission of template. Primers were chosen using Primer Express 2.0 software (Applied Biosystems, Foster City, CA) for optimum use in qPCR. A BLASTN search was performed in GenBank to ensure that all primers were unique to the gene of interest. To avoid amplification from genomic DNA contamination, all primer sets spanned a large exon-intron-exon junction (see Table 1). All qPCR reactions were performed in the ABI prism 7700 (Applied Biosystems) in triplicate with 12.5 µl SYBR Green Master Mix (Applied Biosystems), 5 µl of 5x cDNA, and primers in a total volume of 25 µl. Initial denaturation occurred at 95°C for 10 min, followed by 40 cycles of 15 s at 95°C and 15 s at 60°C. A heat dissociation was performed at the end of every run to assure specificity. Optimum primer concentration was determined for each primer pair by comparing the {Delta}Rn from a matrix of forward and reverse primer concentrations to the Ct (the fractional cycle number at which the fluorescence passes the threshold) for each pair. Optimum primer concentrations are listed in Table 2. An arbitrary threshold of fluorescence was set within the exponential phase of amplification for each gene product and kept constant for all assays. The cycle at which amplification of the product exceeded this threshold was determined and designated as the Ct value. Dissociation curves, gel analysis, and sequencing of certain PCR products confirmed gene-specific product amplification.


View this table:
[in this window]
[in a new window]
  		TABLE 2. Quantitative real-time PCR assay parameters and performance analysis

Standard Curves and Calculations

Relative standard curves were created with serially diluted total RNA samples from the ovaries and SCN used in the experiments (modified from [41]). Three sets of pooled RNA samples were serially diluted by a factor of two starting at 2 µg and ending at 125 ng. This range was determined to be well within the detection level and sensitivity of the qPCR assay. RNA samples used for standard curves are representative of 36 animals. After reverse transcription, the resulting cDNA was PCR amplified in the ABI 7700 in duplicate. Average Ct values were plotted on a logarithmic scale against the log2 of the total RNA concentration. All amplicons had a good doubling efficiency and demonstrated a linear relationship with total RNA concentration (Table 2). Resulting standard curves were used to calculate the relative amount of Arntl and Per2 in the ovary. Relative amount was converted to percent of the maximum amount for each experiment. Percentages were averaged, and SEM was calculated for each group. Data are presented as the mean ± SEM.

The standard curve for Arntl was: y = –1.2171x + 33.517 (R2 = 0.9924; E = 1.77); and for Per2, y = –1.1343x + 32.717 (R2 = 0.9924, E = 1.84); where E = efficiency of amplicon doubling with each cycle 2(–1/slope). Resulting standard curves were used to calculate relative amounts of Arntl and Per2 according to the equation: relative amount = 2([Ct – Intercept]/[slope]).

Histological Analysis of mRNA by Digoxigenin-Labeled Probes

Probes The Per2 and Arntl cDNA fragment-containing vectors were linearized with restriction enzymes and then used as templates for sense or antisense cRNA probes. Probes were generated using digoxigenin (DIG)-11-UTP (Roche) with standard protocols for cRNA synthesis. The Per2 cDNA fragment-containing vector was generously donated by Hitoshi Okamura (Kobe University School of Medicine, Kobe, Japan), and the Arntl cDNA fragment-containing vector was generously donated by Alena Sumova (Institute of Physiology, Academy of Sciences of the Czech Republic, Prague, Czech Republic).

In situ hybridization Tissues were fixed, paraffin embedded, and sectioned the same as for IHC. Briefly, after xylenes and hydration in serial ethanol, tissues were treated for 20 min at room temperature in 0.2 N HCl. Sections were treated in 2x saline sodium citrate (SSC; 1x SSC = 150 mM NaCl, 15 mM sodium citrate; pH 7.0) at 70°C, washed in 1x PBS with 0.3% Triton-X (1x PBS = 140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4; pH 7.2), digested in 2 µg/ml Proteinase K (0.1 M Tris buffer, 50 mM EDTA; pH 8.0) for 15 min at 37°C, washed in 1x PBS with 100 mM Glycine, postfixed in 4% paraformaldehyde, treated in 0.25% acetic anhydride in 0.1 M triethanolamine for 10 min, and then washed in 2x SSC. After an incubation step in 4x SSC/50% deionized formamide for 1 h at 42°C, sections were incubated in hybridization solution (Sigma) containing either DIG-labeled Per2 or Arntl cRNA for 16 h at 42°C. Posthybridization rinses were performed in 2x SSC/50% formamide at 42°C, and then with decreasing concentrations of SSC. Sections were prepared for alkaline phosphatase staining by several washes in wash buffer (100 mM Tris-HCl, 150 mM NaCl; pH 7.5) and then in blocking buffer (1% heat-inactivated normal sheep serum in wash buffer). They were then incubated in the alkaline phosphatase-conjugated antibody (Roche) diluted 1:500 in blocking buffer for 2 h at room temperature. After several washes in wash buffer, sections were pretreated for 10 min in detection buffer (100 mM Tris-HCl, 100 mM NaCl, 50 mM MgCl2; pH 9.5). Finally, the sections were incubated in a solution containing 5-bromo-4-chloro-3-indolyl phosphate toluidium salt (0.18 mg/ml) and nitroblue tetrazolium salt (0.45 mg/ml) (Roche) in detection buffer for 30 min. The coloring reaction was stopped by immersing slides in 1xPBS. The use of sense Per2 and Arntl probes revealed no specific hybridization signals in the ovary sections. Additional controls included omission of the cRNA probe, which also revealed no hybridization signals.

Immunoblot

Ovarian and SCN protein was isolated from the final phenol-ethanol fraction using the standard TRIzol protocol (Invitrogen). The SCN sample, used as a positive control for this study, consisted of pooled SCN reduced slices (n = 12) obtained at different times of day. Resuspended protein was quantified using the Micro BCA protein Assay (Pierce, Rockford, IL). Samples were heated at 100°C for 5 min with an equal volume of Laemmli sample buffer/2-mercaptoethanol (Bio-Rad) and then centrifuged at 13 000 x g for 5 min before separation by sodium dodecyl sulfate-PAGE on an 8% gel (25 µg protein/lane). After transfer (30 min at 100 V), nitrocellulose membranes were stained with 1x Ponceau (Sigma) and imaged to assess uniform loading and transfer. Milk (5%; 1 h at room temperature) was used as a blocking agent. Blots were incubated with primary antibody solution overnight at 4°C (PER2 affinity purified 5 µg/ml; ADI, San Antonio, TX; ARNTL affinity purified 5 µg/ml; ABR, Golden, CO). Blots were washed and incubated in secondary antibody (donkey anti-rabbit:horseradish peroxidase [HRP], 1:5 000; 1 h at room temperature). After a final wash, membranes were developed using chemiluminescent substrate according to the manufacturer's protocol (Pierce). Densitometry using the digital imaging software Scion Image for Windows 4.0.3 (Scion Corporation) provided quantitative analysis of data. The relative density of ARNTL and PER2 immunoreactivity was normalized against alpha-tubulin. Data were plotted as fold change from the lowest point.

Immunohistochemistry

Ovaries were fixed in 10% neutral buffered formalin for 48 h at room temperature. After paraffin embedding, 4 µm sections were cut. After deparaffinization, antigen retrieval, and blocking in normal goat serum, slides were incubated overnight in primary antibody, PER21-A (ADI) or ARNTL (ABR). PER2 antibody was diluted to 20 µg/ml and ARNTL antiserum was diluted to 10 µg/ml. Sections were washed and incubated with an HRP-labeled secondary antibody for 1 h (Vector Laboratories, Burlingame, CA). The peroxidase antibody complex was visualized using aminoethyl carbazole substrate (Zymed, San Francisco, CA). Control experiments included omission of the primary antibody and preabsorption with the peptide specific to the primary antibody. Sections were counterstained with hematoxylin.

Statistical Analysis

All statistical analyses were performed in consultation with a statistician (Dr. David Schaeffer, University of Illinois). Least squares ANOVA was used to analyze all data except Arntl data from the cycle-dependence study. Tukey post hoc analysis was used when ANOVA returned a value of P ≤ 0.05. Because the Arntl qPCR data in the cycle-dependence study did not meet the criteria for homogeneity of variance in the least squares ANOVA, these data were analyzed using generalized estimated equations (GEE) model as implemented with SUDAAN version 9.0 (RTI International).

RESULTS

Clock Gene Transcript Expression in the Ovary

Clock gene products known to comprise the core oscillator (Arntl, Clock, Per1, Per2, Cry1), as well as a mammalian ortholog of the Drosophila canonical clock gene, Timeless, were expressed in the rat ovary (Fig. 1). When samples were obtained every 4 h over a 24-h period, distinct patterns of expression emerged for several of the transcripts. Estrous cycle stage was not determined in these animals. Both Arntl (7-fold; P < 0.001) and Per2 (2-fold; P < 0.05) displayed sinusoidal-like expression patterns with highly statistically significant changes over the 24-h sampling period. Ovarian expression of Arntl peaked at ZT 0/24 (Fig. 2A) and was lowest at ZT 12. Ovarian peak expression for Arntl was out of phase with the SCN (Fig. 2A, dotted line); peak expression of Arntl was observed approximately 4 h later in the ovary. Ovarian expression of Per2 peaked at ZT 16, with a nadir at ZT 4 (Fig. 2B). Similar to Arntl, Per2 expression was out of phase with the SCN (dotted line), with the ovarian peak occurring approximately 4–6 h later (Fig. 2B). Arntl and Per2 expression are in complete antiphase to each other in the ovary, which indicates a potential for the molecular clockworks to function within the ovary.


Figure 1
View larger version (54K):
[in this window]
[in a new window]
  		FIG. 1. Clock gene products are expressed in the rat ovary. Representative denaturing polyacrylamide gel analysis of gene-specific qPCR products from rat ovary indicates bands of appropriate sizes for Arntl, Clock, Per1, Per2, Cry1, and Timeless (labeled "Tim") based on primers listed in Table 1


Figure 2
View larger version (28K):
[in this window]
[in a new window]
  		FIG. 2. Quantitative PCR indicates daily cycling of Arntl and Per2 transcripts in the adult rat ovary. Relative amounts of transcript are represented as the percent of maximum expression vs. ZT, where ZT 0 indicates the time of lights-on in the donor colony. Time of lights-off is indicated by the shaded box. Data are mean ± SEM. A) Arntl transcripts cycle in the ovary with peak expression at ZT 24/0 and minimal expression at ZT 12 [38]. Peak expression is out of phase when compared to the SCN. ***Statistical significance (n = 4; P < 0.001) compared to ZT 12 time point, as determined by one-way ANOVA with Tukey post hoc analysis. B) Per2 transcripts cycle in the ovary with peak expression at ZT 16 and minimal expression at ZT 4. Peak expression is out of phase when compared to the SCN. *Statistical significance (n = 4; P < 0.05) compared to ZT 4 time point as determined by one-way ANOVA with Tukey post hoc analysis

To examine which ovarian cell types may contribute to oscillatory expression of clock genes, we performed in situ hybridization for Arntl and Per2 in the ovary. ZT 0 and ZT 15 were selected to represent those times when the two transcripts were expected to be high and low, as indicated by qPCR data (Fig. 2). Estrous cycle stage was not assessed in these animals. Arntl transcripts were present in granulosa and theca layers of growing and antral follicles, as well as corpora lutea and stromal fibroblasts (Fig 3, A–D). When comparisons were made between ZT 0 and ZT 15, it was apparent that Arntl transcripts were elevated in granulosa cells, and particularly in theca layers of growing and antral follicles at ZT 0. Levels of Arntl transcripts did not change in corpora lutea or in the ovarian stroma. Similarly, Per2 transcripts were present in granulosa and theca layers of both growing and antral follicles (Fig. 3, E–H). Staining for Per2 was also observed in ovarian stroma and corpora lutea (not shown). Per2 transcripts were higher in granulosa and theca cells of growing follicles at ZT 15 (Fig. 3, E and F). Although less apparent, granulosa and theca cells of antral follicles also appeared elevated at ZT 15 (Fig. 3, G and H). Positive controls from rat SCN demonstrate the specificity of the probes. Negative controls, including no probe and sense probe, showed no staining (data not shown). These data support the qPCR results, demonstrating that Arntl and Per2 transcripts are expressed in an out-of-phase relationship in the ovary. Furthermore, these data indicate that Arntl and Per2 are expressed in the same ovarian cell types.


Figure 3
View larger version (87K):
[in this window]
[in a new window]
  		FIG. 3. In situ hybridization for Arntl and Per2 transcripts in the rat ovary at ZT0 and ZT15. A and B) Growing follicles in the rat ovary express the Arntl transcript. Levels appear higher in both granulosa and theca cells at ZT0, but remain unchanged in ovarian stromal fibroblasts. C and D) Arntl transcripts are present in granulosa and theca cells in large antral follicles. Levels appear to be increased at ZT0, but unchanged in corpora lutea. E and F) Per2 transcripts are expressed in ovarian stromal fibroblasts, granulosa and theca cells in growing follicles. Levels are increased at ZT15 in granulosa and theca cells, but remain constant in the stroma. G and H) Per2 transcripts are present in granulosa and theca cells in large antral follicles. Levels appear to be increased at ZT15 in granulosa and theca cells, but unchanged in stroma. I and J) Arntl and Per2 transcript expression in the rat SCN—a positive control. Gr, granulosa; Cl, corpus luteum; arrows point to theca layer; asterisk indicates stroma. Bars = 20 µm

Clock Gene Protein Expression in the Ovary

Immunoblot analysis was performed to determine the protein profile for the clock genes ARNTL and PER2 in the rat ovary. Estrous cycle phase was not determined in these animals. Analysis of protein extracts isolated from the same samples as used for transcript analysis revealed significant 24-h variation in the protein products ARNTL and PER2 (Fig. 4). Peak protein levels for ARNTL (Fig. 4, A and B) were observed at ZT 24/0 (ANOVA, P < 0.01 compared to ZT 12) with minimal levels occurring at ZT 12–16. PER2 levels peaked at ZT 12–16 (ANOVA, P < 0.01 compared to ZT 24/0), with a nadir at ZT 24/0 (Fig. 4, A and C). As previously demonstrated for Arntl and Per2 transcripts, peak expression of ARNTL and PER2 proteins was in complete antiphase.


Figure 4
View larger version (33K):
[in this window]
[in a new window]
  		FIG. 4. A) Representative immunoblot images of adult cycling rat ovarian protein. Oscillating levels of band density for both ARNTL and PER2 immunoreactivity is evident. Alpha tubulin was used as the loading control. Densitometry analysis revealed a 24h oscillation in peak band density with ARNTL (B) peaking at ZT0/24 and PER2 (C) peaking at ZT16 (n = 4). **Statistical significance (P ≤ 0.01, ANOVA with Tukey post hoc analysis) compared to the nadir of expression (ZT16 for ARNTL and ZT24/0 for PER2). Data are means ± SEM

IHC was performed to confirm that cellular localization of ARNTL and PER2 proteins in the rat ovary corresponds to that previously observed for the transcripts (Fig. 3). ZT 4 was selected because both proteins should be detectable at this time at an intermediate level. Estrous cycle phase was not determined in these animals. Immunoreactivity indicated that both proteins were highly expressed in specific ovarian structures (Fig. 5, A and B). Staining was absent or severely reduced upon omission of primary antibody (data not shown) or preabsorption of primary antibody against its peptide (Fig. 5, C and D). ARNTL immunoreactivity was prominent in ovarian stromal fibroblasts (Fig. 5A). However, strong immunoreactivity for ARNTL was also observed in granulosa and theca cells of growing and antral follicles, as well as in corpora lutea. The staining pattern for PER2 was similar. PER2 immunoreactivity was also observed in granulosa and theca cells, and corpora lutea (Fig. 5B). In addition, ARNTL and, especially, PER2 immunoreactivity was strong in oocytes.


Figure 5
View larger version (82K):
[in this window]
[in a new window]
  		FIG. 5. IHC of ARNTL (A and B) and PER2 (C and D) from intact adult rat ovaries. Estrous cycle stage was not determined. Immunoreactivity is red with a blue nuclear hematoxylin counter stain. ARNTL immunolocalization is strong in the ovarian granulosa and thecal cells in growing and antral follicles, as well as stromal cells. PER2 staining is also found in granulosa and theca cells, as well as stromal fibroblasts. B and D) Preabsorption of the primary antibodies with control peptides; immunoreactivity significantly reduced under these conditions, indicating specific immunoreactivity for ARNTL and PER2 antigens in these tissue preparations. Gr, granulosa, arrows point to theca layer, asterisk indicates stroma. Original magnification x20; bars = 10 µm

Estrous Cycle Changes in Clock Gene Expression

Arntl transcript expression patterns were distinctive between two estrous cycle stages. As in Figure 2, Arntl was expressed in a rhythmic pattern, with peak expression occurring near the end of night. On the day of proestrus, Arntl transcript levels were significantly elevated at ZT 18 (ANOVA with Tukey post hoc, P < 0.01) when compared to same time of day on the day of Diestrus I (Fig. 6A). Notably, ZT 18 is approximately 8–10 h after the expected ovulatory LH surge on the day of proestrus. In contrast, Per2 gene expression was not significantly different between the 2 days of the rat estrous cycle, although the pattern of expression on both days displayed a characteristic daily oscillation, with a peak in the first half of the night (Fig. 6B).


Figure 6
View larger version (25K):
[in this window]
[in a new window]
  		FIG. 6. 24 h expression patterns of Arntl and Per2 transcripts across two days of the rat estrous cycle, diestrus I and proestrus, as measured by qPCR. Relative amounts of transcript are represented as the percent of maximum expression vs. ZT. Time of lights-off is indicated by the gray bars. Expression patterns are similar in terms of peak and trough values to those in Figure 1. Data are mean ± SEM. A) Arntl transcripts oscillate during diestrus I and proestrus in the rat ovary. Arntl transcript levels are significantly higher (n = 3; P < 0.01, GEE model) at ZT 18 on the day of proestrus as indicated by **. B) Per2 transcripts oscillate during diestrus I and proestrus in the rat ovary (n = 4–5, P < 0.01, ANOVA). No significant differences were observed in Per2 expression between individual ZTs on these 2 d of the cycle

Effects of Gonadotropins on Clock Gene Expression

Increased expression of Arntl at ZT 18 on the day of proestrus suggests that LH may play a role in regulating expression of this transcript. We used the hypophysectomized, immature rat model to test this hypothesis. Expression patterns and levels of Arntl and Per2 gene products were altered by hCG treatment in juvenile, hypophysectomized rats primed with eCG (control) (Fig. 7). In control animals treated with eCG alone, Arntl transcripts did not change with time of day over the 24 h course of data collection. Arntl displayed highly significant differences between the treatment and control groups across the 24-h time course (P < 0.01, ANOVA). The pattern is very different in the hCG treatment group. As early as 4 h after treatment, Arntl levels show an increasing trend. Arntl transcript levels were significantly elevated compared to the initial time point in the hCG group at 8 and 12 h after treatment (P < 0.05). Similarly, the pattern of Per2 transcript expression was also altered by hCG treatment. Per2 expression in control ovaries (Fig. 7B) showed no evidence of cycling over the 24-h time course of data collection. In contrast, Per2 transcript levels were elevated only 4 h after hCG treatment, and returned to basal levels by 24 h later (ANOVA with Tukey post hoc, P < 0.05). In situ hybridization indicated hCG treatment specifically increases Arntl and Per2 transcripts within thecal layers of large follicles (Fig. 7, C–F). These data strongly indicate that LH can induce cyclic expression of circadian clock genes in the hypophysectomized, immature rat model.


Figure 7
View larger version (54K):
[in this window]
[in a new window]
  		FIG. 7. Effects of hCG on Arntl and Per2 expression patterns in the hypophysectomized, immature rat model as measured by qPCR. All animals were treated with eCG on the first day of the experiment to initiate follicular growth and LH responsiveness. The treatment group received hCG 52 h after eCG treatment. Controls received eCG and then vehicle only. Relative amounts of transcript are represented as the percent of maximum expression vs. time after hCG treatment. Lights off is indicated by gray bars. Data are means ± SEM. A) Expression of Arntl transcripts in the control group does not cycle. Treatment with hCG induces cyclic Arntl expression (n = 4–5; P < 0.05, ANOVA) with a peak occurring at 12 h after treatment (ZT 16) as indicated by *. B) Expression of Per2 transcripts in control also not rhythmic. Treatment with hCG induced cyclic Per2 expression (n = 4–5), with a peak occurring 4 h after hCG treatment (ZT 8; n = 4–5; P < 0.05) as indicated by *. C and D) In situ hybridization for Arntl transcripts indicated an increase in theca cells (arrows) of large follicles at 8 h after treatment. E and F) Human CG also increase Per2 transcripts in theca cells (arrows) at 4 h after treatment. Bars = 10 µm (C–F)

DISCUSSION

Production of fertilizable ova is a meticulously choreographed affair that requires precise timing of events between the ovary and the brain. Coordination of organismic timing is controlled by a circadian timing system, constructed from the dynamic molecular interactions of a core group of "clock" genes. As an initial step toward determining whether a circadian clock has a role in ovarian function, we examined clock gene expression patterns in the rat ovary. The results presented here provide the first report of rhythmic clock gene expression in the ovary, although a few studies have examined clock genes in other components of the female reproductive tract [37, 42]. Arntl and Per2 transcripts and proteins are expressed in appropriate antiphase relationships in the ovary, suggesting that the molecular underpinnings of the clockworks are present. Furthermore, Arntl transcript expression is different on 2 days of the estrous cycle—namely, diestrus I and proestrus. Finally, Arntl and Per2 are upregulated following hCG treatment in juvenile, hypophysectomized animals, suggesting that the circadian-regulated LH surge, required for ovulation, may provide a circadian signal to the ovary.

The ovary expresses each of the genes considered part of the core molecular clock machinery. Similar to other peripheral tissues [18, 43–46] and the SCN, gene products for Arntl and Per2 cycle in the ovary with peaks in expression that are out of phase with each other. In the SCN, increased levels of ARNTL lead to formation of heterodimers with constitutively expressed CLOCK to drive transcription of the Per, Cry, and Tim genes [8]. Accumulating PER and CRY proteins form multimeric complexes that feed back to impede the function of the ARNTL/CLOCK heterodimer, thereby inhibiting their own transcription. In addition, PER proteins act as positive regulators of Arntl transcription [13, 14]. Thus, in the functional mammalian molecular clock first described in the SCN [8], the cycling "positive element," Arntl is expressed in complete antiphase to the "negative elements," Per and Cry; levels of Arntl transcript are elevated when Per and Cry are low, and vice versa. Our data indicate that antiphase expression patterns of positive and negative elements are preserved in the ovary. Furthermore, in accordance with patterns in other peripheral tissues [18, 44, 46], peak expression of these genes in the ovary is shifted by several hours compared to the SCN.

Precise coordination of timing within the hypothalamic-pituitary-ovarian axis is essential to female fertility. Because the circadian clock is fundamental for governance of organismic timing, it seems plausible that female reproductive function may be regulated by central timing mechanisms. Analysis of the circadian influences on female reproductive function has, to date, been restricted to SCN regulation of the LH surge. Required for ovulation, the LH surge occurs on the afternoon of proestrus, when a precisely timed circadian signal manifests concomitant with an appropriate steroidal environment [28, 29]. The circadian nature of this phenomenon is revealed by daily expression of an LH surge in ovariectomized animals with chronic estrogen replacement [28]. Thus, gonadotropin releasing hormone-directed production of LH offers an interesting candidate for circadian regulation of potential ovarian rhythms.

Examination of Arntl and Per2 expression over 2 days of the estrous cycle provides the first evidence that LH may be an important regulator of circadian clock gene rhythms in the ovary. Because our studies were focused on LH as a potential ovarian entrainment factor, we focused on the day of proestrus, and made comparison to another day (diestrus I) when no LH surge occurs. Times of peak expression of Arntl and Per2 were not different between diestrus I and proestrus. However, the Arntl gene product was significantly elevated at ZT 18 on the day of proestrus, approximately 8–10 h after the endogenous LH surge. This finding led to the hypothesis that LH plays a role in regulating expression of this gene product.

We examined this hypothesis directly in the juvenile, hypophysectomized rat. In contrast to the adult, cycling rat, hypophysectomized, juvenile rats, which received eCG to stimulate follicle development and LH responsiveness, showed no evidence of circadian expression patterns for the canonical clock genes, Arntl and Per2. Per2 transcripts were tonically expressed at intermediate levels, whereas, under these same conditions, Arntl transcripts were also arrhythmic, albeit at constitutively low levels. These results are consistent with a lack of circadian clock function, and are reminiscent of the Per2 and Arntl levels observed in the completely arrhythmic mice lacking both Cry1 and Cry2 [13, 47, 48]. Thus, in hypophysectomized, juvenile animals, which have never experienced an LH surge, there is no apparent circadian rhythm in the ovary, and levels of Arntl and Per2 expression mimic those observed in the SCN of known circadian clock mutants. In contrast, hCG treatment induced a circadian pattern of expression for both genes. Furthermore, peak expression of Per2 and Arntl were out of phase with each other. Similar to the estrous cycle study, Arntl was significantly elevated 8–12 h after hCG treatment, suggesting that LH may upregulate expression of the Arntl transcript.

The expression profile for the Per2 transcript was also altered by hCG stimulation in juvenile, hypophysectomized animals. Per2 was elevated 4 h after hCG treatment. Although these results may initially appear contradictory with the findings in the estrous cycle study, it is notable that hCG was administered when Per2 transcript levels were low. In contrast, the endogenous LH surge occurred when Per2 levels were already approaching their maximum; thus, further stimulation of this gene product beyond peak levels may not have been possible. Similarly, in the SCN, light pulses will stimulate Per2 transcripts only when Per2 is endogenously low [49].

Generally, the data from the juvenile, hypophysectomized animals suggest that LH may provide a sufficient signal to initiate clock gene rhythmicity in the ovary. In this model, Per2 was elevated first, followed by an appropriately out-of-phase increase in Arntl after several hours. In the current interlocking feedback loop model of clock gene function, Per2 can drive transcription of Arntl. Whether LH acts through Per2 or a different mechanism to drive ovarian rhythms requires further investigation. Our data highlight the usefulnees of the juvenile, hypophysectomized animal as a model for endocrine induction of circadian rhythmicity in a peripheral tissue.

Induction of Arntl and Per2 transcripts by hCG was particularly evident in the theca layers of growing and antral follicles. Thus, the theca cells may be important for receiving the circadian signal and initiating rhythmicity in developing follicles. If LH is the agent that carries the circadian signal to the ovary, the theca layer is primed to receive this message because LH receptor expression is restricted to the thecal cell compartment in developing follicles [50, 51]. Furthermore, FSH can increase LH receptor expression in granulosa cells [51]. Further experimentation is required to determine the mechanisms underlying increased expression of Per2 and Arntl in response to LH treatment.

Many questions remain regarding the functional significance of circadian clock genes within the ovary. Mice bearing deletions of the negative arm of the circadian transcriptional/translational feedback loop (the Cry and Per genes) have no apparent deficits in reproductive fitness. In contrast, disruption of positive elements (Arntl and Clock) is detrimental to reproduction. Arntl-null mice have irregular estrous cycles, decreased ovulation rates, and fail to maintain pregnancy to term [42]. Clock mutant mice also have impaired fertility [52–54]. Although superficial histological examination of Clock mutant ovaries has revealed no significant differences between numbers of corpora lutea and preovulatory follicles with wild-type tissues [52], comprehensive analysis of the ovary has not been completed in any circadian clock mutant animals. IHC has revealed expression of ARNTL and PER2 in ovarian follicles throughout the latter stages of follicular development and in corpora lutea, as well as in oocytes (Fig. 4). It is clear that successful follicular development requires coordinated communication between the oocyte and the surrounding somatic cells [55]. Thus, it is attractive to hypothesize that clock gene expression may play a role in directing follicular development as the oocyte itself orchestrates the rate of ovarian follicular development by conducting the growth and maturation of the surrounding granulosa and theca layers [56].

Notably, another basic helix-loop-helix, PAS domain protein similar to circadian clock proteins, the aryl hydrocarbon receptor (AhR), is important for normal ovarian follicular development [57]. AhR protein expression patterns in the ovary are similar to ARNTL and PER2 [58]. AhR has been linked to the rate of apoptosis in the developing ovarian germ line, the rate of follicle development, and, ultimately, the rate of ovulation. Likewise, another member of the basic helix-loop-helix (bHLH) proteins, SHARP2, is expressed in granulosa and theca interna layers of the developing follicles and is also stimulated by hCG [59]. It is not unreasonable to speculate that clock genes could be involved in follicular growth and maturation through potential interactions with AhR, SHARP2, and other bHLH containing transcription factors within the ovarian follicle. Finally, interactions between clock genes and the cell cycle may be reflected in expression of these genes in granulosa and theca cells of growing follicles [60–63]. The results of this study suggest that clock genes have the potential to influence ovarian follicular function, or, at the very least, may assure maintenance of the delicate balance of timing within the female reproductive axis.

Physiologic circadian rhythmicity has long been considered dependent upon the pacemaker activity of the master clock located in the SCN [3, 4, 7], although this concept has recently been the subject of debate [26, 64]. SCN lesion disrupts behavioral and endocrine circadian rhythms [65, 66], as well as oscillations in peripheral clocks [22, 25]. Transplant studies indicate that the SCN determines the circadian period in the host [67]. We hypothesize that an SCN-derived LH surge provides timing cues to the ovary. Investigation of the effects of LH on other reproductive tissues, as well as nonreproductive tissues, may provide additional insight into the effects of LH on clock gene expression. However, alternate timing cues should also be considered. Transneuronal viral-tracing techniques have revealed that the SCN has direct neuronal connections with the liver and pancreas through the autonomic nervous system [68]. Circadian timing cues could also reach the ovary through direct neuronal connections from the SCN via the autonomic nervous system [69]. Continuing research efforts are aimed at determining the role of circadian clock genes in the ovary, as well as exploring whether the SCN ultimately provides timing cues for synchronization of ovarian function within the female reproductive axis.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Primers used for qPCR analysis and amplicon characteristics

ACKNOWLEDGMENTS

The authors are grateful to M. Nakai for technical assistance with IHC, D. Schaeffer for assistance with statistical analysis, J. Hickok for development of qPCR techniques, and J. Jorgensen and K. Bottum for comments on the manuscript.

FOOTNOTES

1 Supported by National Institutes of Health grant ES012948 and the Illinois Governor's Venture Technology Fund (to S.A.T.). Back

2 Correspondence: Shelley A. Tischkau, Department of Veterinary Biosciences, University of Illinois at Urbana-Champaign, 3840 VMBSB, 2001 S. Lincoln Ave., Urbana, IL 61802. FAX: 217-244-1652; tischkau{at}uiuc.edu Back

Received: 4 January 2006.

First decision: 27 January 2006.

Accepted: 26 June 2006.

REFERENCES

  1. de Vries MJ, Nunes Cardozo B, van der Want J, de Wolf A, Meijer JH, Glutamate immunoreactivity in terminals of the retinohypothalamic tract of the brown Norwegian rat. Brain Res 1993 612:231-237[CrossRef][Medline]
  2. Ding JM, Chen D, Weber ET, Faiman LE, Rea MA, Gillette MU, Resetting the biological clock: mediation of nocturnal circadian shifts by glutamate and NO. Science 1994 266:1713-1717[Abstract/Free Full Text]
  3. Ralph MR, Foster RG, Davis FC, Menaker M, Transplanted suprachiasmatic nucleus determines circadian period. Science 1990 247:975-978[Abstract/Free Full Text]
  4. Reppert SM, Weaver DR, Coordination of circadian timing in mammals. Nature 2002 418:935-941[CrossRef][Medline]
  5. Shibata S, Watanabe A, Hamada T, Ono M, Watanabe S, N-methyl-D-aspartate induces phase shifts in circadian rhythm of neuronal activity of rat SCN in vitro. Am J Physiol 1994 267:R360-R364[Medline]
  6. Shirakawa T, Moore RY, Glutamate shifts the phase of the circadian neuronal firing rhythm in the rat suprachiasmatic nucleus in vitro. Neurosci Lett 1994 178:47-50[CrossRef][Medline]
  7. Weaver DR, The suprachiasmatic nucleus: a 25-year retrospective. J Biol Rhythms 1998 13:100-112[Abstract]
  8. Okamura H, Yamaguchi S, Yagita K, Molecular machinery of the circadian clock in mammals. Cell Tissue Res 2002 309:47-56[CrossRef][Medline]
  9. Hastings MH, Herzog ED, Clock genes, oscillators, and cellular networks in the suprachiasmatic nuclei. J Biol Rhythms 2004 19:400-413[Abstract]
  10. King DP, Takahashi JS, Molecular genetics of circadian rhythms in mammals. Annu Rev Neurosci 2000 23:713-742[CrossRef][Medline]
  11. Okamura H, Integration of mammalian circadian clock signals: from molecule to behavior. J Endocrinol 2003 177:3-6[Abstract]
  12. Harms E, Kivimae S, Young MW, Saez L, Posttranscriptional and posttranslational regulation of clock genes. J Biol Rhythms 2004 19:361-373[Abstract]
  13. Shearman L, Sriram S, Weaver D, Maywood ES, Chaves I, Zheng B, Kume K, Lee C, van der Horst G, Hastings M, Reppert S, Interacting molecular loops in the mammalian circadian clock. Science 2000 288:1013-1019[Abstract/Free Full Text]
  14. Preitner N, Damiola F, Lopez-Molina L, Zakany J, Duboule D, Albrecht U, Schibler U, The orphan nuclear receptor REV-ERBalpha controls circadian transcription within the positive limb of the mammalian circadian oscillator. Cell 2002 110:251-260[CrossRef][Medline]
  15. Delaunay F, Laudet V, Circadian clock and microarrays: mammalian genome gets rhythm. Trends Genet 2002 18:595-597[CrossRef][Medline]
  16. Albrecht U, Sun ZS, Eichele G, Lee CC, A differential response of two putative mammalians circadian regulators, mper1 and mper2, to light. Cell 1997 91:1055-1064[CrossRef][Medline]
  17. Sun ZS, Albrecht U, Zhuchenko O, Bailey J, Eichele G, Lee CC, RIGUI, a putative mammalian ortholog of the Drosophila period gene. Cell 1997 90:1003-1011[CrossRef][Medline]
  18. Zylka MJ, Shearman LP, Weaver DR, Reppert SM, Three period homologs in mammals: differential light response in the suprachiasmatic circadian clock and oscillating transcripts outside the brain. Neuron 1998 20:1103-1110[CrossRef][Medline]
  19. Oishi K, Sakamoto K, Okada T, Nagase T, Ishida N, Humoral signals mediate the circadian expression of rat period homologue (rPer2) mRNA in peripheral tissues. Neurosci Lett 1998 256:117-119[CrossRef][Medline]
  20. Balsalobre A, Damiola F, Schibler U, A serum shock induces circadian gene expression in mammalian tissue culture cells. Cell 1998 93:929-937[CrossRef][Medline]
  21. Balsalobre A, Brown SA, Marcacci L, Tronche F, Kellendonk C, Reichardt HM, Schutz G, Schibler U, Resetting of circadian time in peripheral tissues by glucocorticoid signaling. Science 2000 289:2344-2347[Abstract/Free Full Text]
  22. Sakamoto K, Nagase T, Fukui H, Horikawa K, Okada T, Tanaka H, Sato K, Miyake Y, Ohara O, Kako K, Ishida N, Multitissue circadian expression of rat period homolog (rPer2) mRNA is governed by the mammalian circadian clock, the suprachiasmatic nucleus in the brain. J Biol Chem 1998 273:27039-27042[Abstract/Free Full Text]
  23. Durgan DJ, Hotze MA, Tomlin TM, Egbejimi OA, Graveleau C, Abel ED, Shaw CA, Bray MS, Hardin PE, Young ME, The intrinsic circadian clock with the cardiomyocyte. Am J Physiol Heart Circ Physiol 2005 289:H1530-H1541[Abstract/Free Full Text]
  24. Oishi K, Miyazaki K, Ishida N, Functional CLOCK is not involved in the entrainment of peripheral clocks to the restricted feeding: entrainable expression of mPer2 and BMAL1 mRNAs in the heart of Clock mutant mice on Jcl:ICR background. Biochem Biophys Res Commun 2002 298:198-202[CrossRef][Medline]
  25. Akhtar RA, Reddy AB, Maywood ES, Clayton JD, King VM, Smith AG, Gant TW, Hastings MH, Kyriacou CP, Circadian cycling of the mouse liver transcriptome, as revealed by cDNA microarray, is driven by the suprachiasmatic nucleus. Curr Biol 2002 12:540-550[CrossRef][Medline]
  26. Yoo SH, Yamazaki S, Lowrey PL, Shimomura K, Ko CH, Buhr ED, Siepka SM, Hong HK, Oh WJ, Yoo OJ, Menaker M, Takahashi JS, PERIOD2::LUCIFERASE real-time reporting of circadian dynamics reveals persistent circadian oscillations in mouse peripheral tissues. Proc Natl Acad Sci U S A 2004 101:5339-5346[Abstract/Free Full Text]
  27. Wiegand SJ, Terasawa E, Discrete lesions reveal functional heterogeneity of suprachiasmatic structures in regulation of gonadotropin secretion in the female rat. Neuroendocrinology 1982 34:395-404[CrossRef][Medline]
  28. Legan SJ, Coon GA, Karsch FJ, Role of estrogen as initiator of daily surges in the ovariectomized rat. Endocrinology 1975 96:50-56[Medline]
  29. Legan SJ, Karsch FJ, A daily signal for the LH surge in the rat. Endocrinology 1975 96:57-62[Abstract]
  30. Turek FW, Losee-Olson S, Swann JM, Horwath K, Van Cauter E, Milette JJ, Circadian and seasonal control of neuroendocrine-gonadal activity. J Steroid Biochem 1987 27:573-579[CrossRef][Medline]
  31. Alvarez JD, Sehgal A, The thymus is similar to the testis in its pattern of circadian clock gene expression. J Biol Rhythms 2005 20:111-121[Abstract]
  32. Alvarez JD, Chen D, Storer E, Sehgal A, Non-cyclic and developmental stage-specific expression of circadian clock proteins during murine spermatogenesis. Biol Reprod 2003 69:81-91[Abstract/Free Full Text]
  33. Morse D, Cermakian N, Brancorsini S, Parvinen M, Sassone-Corsi P, No circadian rhythms in testis: Period1 expression is clock independent and developmentally regulated in the mouse. Mol Endocrinol 2003 17:141-151[Abstract/Free Full Text]
  34. Miyamoto Y, Sancar A, Circadian regulation of cryptochrome genes in the mouse. Brain Res Mol Brain Res 1999 71:238-243[Medline]
  35. Bittman EL, Doherty L, Huang L, Paroskie A, Period gene expression in mouse endocrine tissues. Am J Physiol Regul Integr Comp Physiol 2003 285:R561-R569[Abstract/Free Full Text]
  36. Tong Y, Guo H, Brewer JM, Lee H, Lehman MN, Bittman EL, Expression of haPer1 and haBmal1 in Syrian hamsters: heterogeneity of transcripts and oscillations in the periphery. J Biol Rhythms 2004 19:113-125[Abstract]
  37. Kennaway DJ, Varcoe TJ, Mau VJ, Rhythmic expression of clock and clock-controlled genes in the rat oviduct. Mol Hum Reprod 2003 9:503-507[Abstract/Free Full Text]
  38. Tischkau SA, Mitchell JW, Pace LA, Barnes JW, Barnes JA, Gillette MU, Protein kinase G type II is required for night-to-day progression of the mammalian circadian clock. Neuron 2004 43:539-549[CrossRef][Medline]
  39. Gillette MU, Reppert SM, The hypothalamic suprachiasmatic nuclei: circadian patterns of vasopressin secretion and neuronal activity in vitro. Brain Res Bull 1987 19:135-139[CrossRef][Medline]
  40. Petroff BK, Gao X, Rozman KK, Terranova PF, Interaction of estradiol and 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in an ovulation model: evidence for systemic potentiaion and local ovarian effects. Reproductive Toxicology 2000 14:247-255[CrossRef][Medline]
  41. Bustin SA, Absolute quantification of mRNA using real-time reverse transcription polymerase chain reaction assays. J Mol Endocrinol 2000 25:169-193[Abstract]
  42. Kennaway DJ, The role of circadian rhythmicity in reproduction. Hum Reprod Update 2005 11:91-101[Abstract/Free Full Text]
  43. Oishi K, Fukui H, Ishida N, Rhythmic expression of BMAL1 mRNA is altered in Clock mutant mice: differential regulation in the suprachiasmatic nucleus and peripheral tissues. Biochem Biophys Res Commun 2000 268:164-171[CrossRef][Medline]
  44. Ando H, Yanagihara H, Hayashi Y, Obi Y, Tsuruoka S, Takamura T, Kaneko S, Fujimura A, Rhythmic mRNA expression of clock genes and adipocytokines in mouse visceral adipose tissue. Endocrinology 2005 146:5631-5636[Abstract/Free Full Text]
  45. Yamamoto T, Nakahata Y, Soma H, Akashi M, Mamine T, Takumi T, Transcriptional oscillation of canonical clock genes in mouse peripheral tissues. BMC Molecular Biology 2004 5:18[CrossRef][Medline]
  46. Muhlbauer E, Wolgast S, Finckh U, Peschke D, Peschke E, Indication of circadian oscillations in the rat pancreas. FEBS Lett 2004 564:91-96[CrossRef][Medline]
  47. Okamura H, Miyake S, Sumi Y, Yamaguchi S, Yasui A, Muijtjens M, Hoeijmakers J, van der Horst G, Photic induction of mPer1 and mPer2 in Cry-deficient mice lacking a biological clock. Science 1999 286:2531-2534[Abstract/Free Full Text]
  48. Vitaterna MH, Selby CP, Todo T, Niwa H, Thompson C, Fruechte EM, Hitomi K, Thresher RJ, Ishikawa T, Miyazaki J, Takahashi JS, Sancar A, Differential regulation of mammalian period genes and circadian rhythmicity by cryptochromes 1 and 2. Proc Natl Acad Sci U S A 1999 96:12114-12119[Abstract/Free Full Text]
  49. Shearman LP, Zylka MJ, Weaver DR, Kolakowski LF, Jr, Reppert SM, Two period homologs: circadian expression and photic regulation in the suprachiasmatic nuclei. Neuron 1997 19:1261-1269[CrossRef][Medline]
  50. Bao B, Kumar N, Karp R, Garverick H, Sundaram K, Estrogen receptor-beta expression in relation to the expression of luteinizing hormone receptor and cytochrome P450 enzymes in rat ovarian follicles. Biol Reprod 2000 63:1747-1755[Abstract/Free Full Text]
  51. Hillier SG, Gonadotropic control of ovarian follicular growth and development. Mol Cell Endocrinol 2001 179:39-46[CrossRef][Medline]
  52. Miller BH, Olson SL, Turek FW, Levine JE, Horton TH, Takahashi JS, Circadian clock mutation disrupts estrous cyclicity and maintenance of pregnancy. Curr Biol 2004 14:1367-1373[CrossRef][Medline]
  53. Kennaway DJ, Boden MJ, Voultsios A, Reproductive performance in female Clock(Delta19) mutant mice. Reprod Fertil Dev 2005 16:801-810[CrossRef][Medline]
  54. Dolatshad H, Campbell EA, O'Hara L, Maywood ES, Hastings MH, Johnson MH, Developmental and reproductive performance in circadian mutant mice. Hum Reprod 2006 21:68-79[Abstract/Free Full Text]
  55. Eppig JJ, Oocyte control of ovarian follicular development and function in mammals. Reproduction 2001 122:829-838[Abstract]
  56. Eppig JJ, Wigglesworth K, Pendola FL, The mammalian oocyte orchestrates the rate of ovarian follicular development. Proc Natl Acad Sci U S A 2002 99:2890-2894[Abstract/Free Full Text]
  57. Benedict JC, Miller KP, Lin TM, Greenfeld C, Babus JK, Peterson RE, Flaws JA, Aryl hydrocarbon receptor regulates growth, but not atresia, of mouse preantral and antral follicles. Biol Reprod 2003 68:1511-1517[Abstract/Free Full Text]
  58. Robles R, Morita Y, Mann KK, Perez GI, Yang S, Matikainen T, Sherr DH, Tilly JL, The aryl hydrocarbon receptor, a basic helix-loop-helix transcription factor of the PAS gene family, is required for normal ovarian germ cell dynamics in the mouse. Endocrinology 2000 141:450-453[Abstract/Free Full Text]
  59. Yamada K, Kawata H, Mizutani T, Arima T, Yazawa T, Matsuura K, Shou Z, Sekiguchi T, Yoshino M, Kajitani T, Miyamoto K, Gene expression of basic helix-loop-helix transcription factor, SHARP-2, is regulated by gonadotropins in the rat ovary and MA-10 cells. Biol Reprod 2004 70:76-82[Abstract/Free Full Text]
  60. Grundschober C, Delaunay F, Puhlhofer A, Triqueneaux G, Laudet V, Bartfai T, Nef P, Circadian regulation of diverse gene products revealed by mRNA expression profiling of synchronized fibroblasts. J Biol Chem 2001 276:46751-46758[Abstract/Free Full Text]
  61. Granda TG, Liu XH, Smaaland R, Cermakian N, Filipski E, Sassone-Corsi P, Levi F, Circadian regulation of cell cycle and apoptosis proteins in mouse bone marrow and tumor. Faseb J 2005 19:304-306[Abstract/Free Full Text]
  62. Oishi K, Miyazaki K, Kadota K, Kikuno R, Nagase T, Atsumi G, Ohkura N, Azama T, Mesaki M, Yukimasa S, Kobayashi H, Iitaka C, et al Genome-wide expression analysis of mouse liver reveals CLOCK-regulated circadian output genes. J Biol Chem 2003 278:41519-41527[Abstract/Free Full Text]
  63. Lowrey PL, Takahashi JS, Mammalian circadian biology: elucidating genome-wide levels of temporal organization. Annu Rev Genomics Hum Genet 2004 5:407-441[CrossRef][Medline]
  64. Welsh DK, Yoo SH, Liu AC, Takahashi JS, Kay SA, Bioluminescence imaging of individual fibroblasts reveals persistent, independently phased circadian rhythms of clock gene expression. Curr Biol 2004 14:2289-2295[CrossRef][Medline]
  65. Stephan FK, Zucker I, Circadian rhythms in drinking bevavior and locomoter activity of rats are eliminated by hypothalamic lesions. Proc Natl Acad Sci U S A 1972 69:1583-1586[Abstract/Free Full Text]
  66. Moore RY, Eichler VB, Loss of a circadian adrenal corticosterone rhythm following suprachiasmatic lesions in the rat. Brain Res 1972 42:201-206[CrossRef][Medline]
  67. LeSauter J, Lehman MN, Silver R, Restoration of circadian rhythmicity by transplants of SCN "micropunches.". J Biol Rhythms 1996 11:163-171[Abstract/Free Full Text]
  68. Kalsbeek A, La Fleur S, Van Heijningen C, Buijs RM, Suprachiasmatic GABAergic inputs to the paraventricular nucleus control plasma glucose concentrations in the rat via sympathetic innervation of the liver. J Neurosci 2004 24:7604-7613[Abstract/Free Full Text]
  69. Buijs RM, Hermes MH, Kalsbeek A, The suprachiasmatic nucleus-paraventricular nucleus interactions: a bridge to the neuroendocrine and autonomic nervous system. Prog Brain Res 1998 119:365-382[Medline]



This article has been cited by other articles:


Home page
 Toxicol SciHome page
M. E. Andersen, H. J. Clewell III, E. Bermudez, G. A. Willson, and R. S. Thomas
Genomic Signatures and Dose-Dependent Transitions in Nasal Epithelial Responses to Inhaled Formaldehyde in the Rat
Toxicol. Sci., October 1, 2008; 105(2): 368 - 383.
[Abstract] [Full Text] [PDF]

Home page
 J Biol RhythmsHome page
M. Mukai, T.-M. Lin, R. E. Peterson, P. S. Cooke, and S. A. Tischkau
Behavioral Rhythmicity of Mice Lacking AhR and Attenuation of Light-Induced Phase Shift by 2,3,7,8-Tetrachlorodibenzo-p-Dioxin
J Biol Rhythms, June 1, 2008; 23(3): 200 - 210.
[Abstract] [PDF]

Home page
 EndocrinologyHome page
N. Nakao, S. Yasuo, A. Nishimura, T. Yamamura, T. Watanabe, T. Anraku, T. Okano, Y. Fukada, P. J. Sharp, S. Ebihara, et al.
Circadian Clock Gene Regulation of Steroidogenic Acute Regulatory Protein Gene Expression in Preovulatory Ovarian Follicles
Endocrinology, July 1, 2007; 148(7): 3031 - 3038.
[Abstract] [Full Text] [PDF]

Home page
 J EndocrinolHome page
P.-J. He, M. Hirata, N. Yamauchi, S. Hashimoto, and M.-a. Hattori
The disruption of circadian clockwork in differentiating cells from rat reproductive tissues as identified by in vitro real-time monitoring system
J. Endocrinol., June 1, 2007; 193(3): 413 - 420.
[Abstract] [Full Text] [PDF]

Home page
 Toxicol SciHome page
M. Mukai and S. A. Tischkau
Effects of Tryptophan Photoproducts in the Circadian Timing System: Searching for a Physiological Role for Aryl Hydrocarbon Receptor
Toxicol. Sci., January 1, 2007; 95(1): 172 - 181.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
75/4/624    most recent
biolreprod.106.050732v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted