Leptin and Leptin Receptor Expression in the Rat Ovary

0013-7227/03/$15.00/0 Printed in U.S.A.

Endocrinology 144(11):5006 –5013 Copyright © 2003 by The Endocrine Society doi: 10.1210/en.2003-0584

Leptin and Leptin Receptor Expression in the Rat Ovary NATALIE K. RYAN, KYLIE H. VAN DER HOEK, SARAH A. ROBERTSON,

AND

ROBERT J. NORMAN

Reproductive Medicine Unit, Department of Obstetrics and Gynecology, The University of Adelaide, The Queen Elizabeth Hospital, Woodville, South Australia 5011, Australia Leptin is an important satiety hormone and reproductive regulator and is found, along with its receptors, throughout the ovary. To date, the changes in ovarian expression of both of these proteins throughout the estrous cycle has not been studied, and the examination of protein expression has not distinguished between different forms of the receptor. In this study leptin mRNA expression in the immature gonadotropinprimed rat ovary increased 3-fold after human chorionic gonadotropin administration, followed by a dramatic increase in mRNA for both the short form (Ob-Ra) and the long form (Ob-Rb) of the leptin receptor (⬃8- and 7-fold, respectively). A

L

EPTIN, THE PRODUCT of the obese (ob) gene, is an important satiety and reproductive hormone classically secreted by adipose tissue. It has been discovered to be synthesized in many other organs, including the stomach, placenta, and ovary (1– 4). Leptin is essential in maintaining normal reproductive function, as mice deficient in leptin (ob/ob) are not only obese, but are also infertile (1). The administration of leptin to these mice can restore normal weight and rescue reproductive function, whereas weight loss alone is unable to reinstate fertility (5). Leptin has significant direct effects on ovarian granulosa and thecal cells. In vitro experiments have shown leptin stimulation of bovine insulininduced thecal cell proliferation and inhibition of insulininduced progesterone and estradiol production by granulosa cells and insulin-induced progesterone and androstenedione production by thecal cells (6, 7). Leptin also inhibits IGF-I augmentation of steroidogenesis in human granulosa and thecal cells and rat granulosa cells (8, 9) and can prevent granulosa cell apoptosis in rat ovaries (10). There is also evidence of adsorption of serum-borne leptin into ovarian tissues, as administration of exogenous radioactive leptin results in accumulation in the ovary (10). We have previously shown that macrophages, abundant in the stroma and theca at the time of ovulation (11), are critical for normal follicular rupture (12). Leptin acts to enhance cytokine production (particularly granulocyte-macrophage and granulocyte colony-stimulating factor) and phagocytosis in murine peritoneal macrophages (13). Leptin can also stimulate proliferation and activation as well as cytokine and receptor antagonist production (TNF␣, IL-6, and IL-1 receptor agonist) in circulating human monocytes (14, 15). The phagocytic function of peritoneal macrophages from both ob/ob and db/db (deficient in functional leptin reAbbreviations: CT, Critical threshold; eCG, equine chorionic gonadotropin; hCG, human chorionic gonadotropin; LHR, LH receptor; NGS, normal goat serum; STAT, signal transducer and activator of transcription.

corresponding increase in mRNA expression of the receptor was not observed in isolated preovulatory follicles. Using immunohistochemistry, we observed protein expression of the long form of the leptin receptor (Ob-Rb) in the ovary, with high intensities observed in oocytes and endothelial cells as well as thecal cells and corpora lutea. These results suggest that ovarian expression of leptin and its receptor is regulated across the cycle by gonadotropins, with peak expression at ovulation, indicating a possible involvement in oocyte maturation, angiogenesis, follicle rupture, or subsequent corpus luteum formation. (Endocrinology 144: 5006 –5013, 2003)

ceptors) mice is decreased, and administration of leptin to macrophages from ob/ob, but not db/db, mice can return phagocytic function to normal, indicating that macrophages use functional, long-form leptin receptors for leptin responsiveness (16). These studies all indicate that leptin may be a critical player in the regulation of leukocytes in the inflammatory-like reactions in the ovary that are intimately associated with ovulation and corpus luteum formation. The leptin receptor (Ob-R) is a transmembrane receptor found in many tissues, including the hypothalamus, lung, kidney, and many cells of the ovary, including thecal cells, granulosa cells, and oocytes (17–19). There are six known splice variants of the leptin receptor, all with the same extracellular domain, but with differing intracellular domains (Ob-Ra to Ob-Rf). The Ob-Rb isoform contains a 303-amino acid cytoplasmic domain and is considered to be the only splice variant capable of signal transduction across the cell membrane via signal transducer and activator of transcription-3 (STAT3), which leads to altered gene transcription (20, 21). Recently, however, it has been proposed that the mitogen-activated pathway may be a more important signal transducer in reproductive tissues, as leptin in placental cells does not activate the STAT pathway (22). Others have shown that mice with a disrupted Ob-Rb-STAT3 signal are obese, yet fertile, unlike db/db mice, which are obese and infertile (23, 24). One of the short forms of the leptin receptor, Ob-Ra, has been proposed to play a role in leptin transport, particularly in the brain, where high levels of Ob-Ra are detected (25). Ob-Ra mRNA levels fluctuate in the rat placenta, where it is suggested that this receptor acts to transport leptin to the fetus during development (26). We have previously shown that there are high levels of leptin receptor protein in the thecal cell layer of preovulatory follicles in the mouse and that treatment of mice with exogenous leptin increases the rate of oocyte maturation in in vitro cultured preovulatory follicles (27). We have also shown in the rat that in vivo and in vitro administration of leptin to

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rats decreases their ovulation rate (28) and that ovarian Ob-Rb mRNA receptor levels in the natural rat estrous cycle increase at metestrus (29). As leptin has significant effects on ovarian function, and the expression of the ligand and its receptor appear to be fundamental to the understanding of how this protein acts, it is important to further clarify the importance of gonadotropins in regulating the expression of these two proteins in the ovary. Our aim was to describe the temporal pattern of leptin and leptin receptor expression across the gonadotropin-primed cycle in the ovaries of immature rats. This model was used to avoid the confounding effects of the presence of corpora lutea from previous cycles. We also set out to identify the cellular localization of Ob-Rb expression in ovaries by immunohistochemistry, rather than all forms of leptin receptor as has been done previously, and to evaluate the importance of follicular vs. stromal tissues in contributing to cycle-related changes. Materials and Methods Animals Sexually immature Sprague Dawley rats were housed at 24 C on a 14-h light, 10-h dark cycle with water and pelleted food available ad libitum. Approval for the experiments was obtained from the animal ethics committees of both The Queen Elizabeth Hospital and The University of Adelaide, and animals were handled in accordance with the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes. Rats were killed either at 28 d of age (immature) or at various times after ip injection at 26 d of age with 10 IU equine chorionic gonadotropin (eCG; Intervet, Boxmeer, The Netherlands), followed 48 h later by 10 IU human chorionic gonadotropin (hCG; Pregnyl, Organon, Oss, The Netherlands).

Tissue and plasma collection for RNA isolation Rats were killed by anesthesia [400 ␮l of a 1:1 mixture of ketamil (100 mg/ml ketamine, Troy Laboratories Pty. Ltd., Smithfield, Australia) and Rompun (20 mg/ml xylazine, Bayer AG, Leverkusen, Germany)], followed by cardiac puncture to collect blood and then cervical dislocation. Ovaries were removed aseptically, cleaned of surrounding bursa along with its attached fat, placed into Eppendorf tubes, and snap-frozen in liquid nitrogen. The blood obtained was collected into EDTA-coated tubes (Greiner Bio-one GmbH, Kremsmu¨ nster, Austria) and centrifuged to collect plasma, which was frozen at ⫺80 C for future assays. Five or six animals were killed at each time point, resulting in 10 –12 ovaries/group.

Follicle culture For collection of preovulatory follicles, 26-d-old immature rats were primed with 10 IU eCG. Forty-eight hours later, ovaries were removed,

and large, preovulatory follicles (⬎550 ␮m in diameter) were dissected from the ovaries under a dissecting microscope and collected in HEPESbuffered tissue culture medium (ICN Biochemicals Inc., Costa Mesa, CA) supplemented with 0.3% BSA. Follicles were then either lysed and the RNA was extracted immediately or were transferred to stoppered glass Vacutainer tubes (BD Biosciences, Franklin Lakes, NJ) in 1 ml ␣MEM (Trace Biosciences Pty. Ltd., Victoria, Australia) supplemented with 25 mm sodium bicarbonate (Sigma-Aldrich Corp., St. Louis, MO) and 5% heat-inactivated normal rat serum with or without 10, 100, or 1000 mIU/ml hCG. The tubes were then gassed with 95% O2/5% CO2 and placed in a 37 C shaking water bath for 9 h. Culture medium was removed and frozen at ⫺20 C for steroid assays, follicles were removed, and cells were lysed with 350 ␮l lysis buffer (Qiagen, Valencia, CA), then stored at ⫺80 C for future RNA isolation. Additionally some preovulatory follicles were dissected from ovaries 9 h after administration of 10 IU hCG in vivo. Five or six follicles were used in each extraction, and each experiment was replicated five times.

RNA isolation and measurement RNA was isolated from whole frozen ovaries and frozen follicles using an RNeasy kit (Qiagen) and was also treated using deoxyribonuclease I (Qiagen) to remove any residual DNA. The RNA was then dissolved in 50 ␮l (whole ovaries) or 20 ␮l (follicles) ribonuclease-free water and frozen at ⫺80 C. RNA yields were quantified by either spectrophotometry measurements at 260 nm or the more sensitive Ribogreen RNA quantitation kit (Molecular Probes, Eugene, OR) according to the manufacturer’s instructions.

Quantitative RT-PCR For each sample, 2 ␮g RNA for whole ovaries and 1 ␮g RNA for follicles were diluted in PCR-grade water and reverse transcribed using random primers (Roche, Castle Hill, Australia) and a Superscript II RT kit (Life Technologies, Inc., Grand Island, NY) according to the manufacturer’s instructions. Negative controls omitting RNA and/or Superscript II enzyme were included.

Primers Specific primers for rat ␤-actin, leptin (Ob), and leptin receptor (ObRb and Ob-Ra forms) and mouse LH receptor (LHR) were designed on Primer Express software (PE Applied Biosystems, Foster City, CA), manufactured by Geneworks (Adelaide, Australia; Table 1). Both ␤-actin and Ob-Ra products crossed intron/exon boundaries, and all RT-PCR products were validated by sequencing. Samples were run in triplicate on an ABI GeneAmp 5700 sequence detection system. They were subjected to 40 cycles of amplification at 95 C for 15 sec, followed by 60 C for 60 sec using 3 ␮l diluted cDNA (1:30), 10 ␮l SYBR green buffer (PE Applied Biosystems), and 10 pmol of each primer. Negative controls omitting template cDNA were included in each run. After RT-PCR amplification, a dissociation analysis was run on the products to ensure that one product only was produced during

TABLE 1. Primer sets used for quantitative RT-PCR Name

Oligonucleotide sequences

Product length (bp)

Genbank accession no.

Rat ␤-actin

Sense 5⬘-CCGTAAAGACCTCTATGCCAACA-3⬘ Antisense 5⬘-GCTAGGAGCCAGGGCAGTAAT-3⬘

103

gi: 55574

Rat leptin (Ob)

Sense 5⬘-TTTCACACACGCAGTCGGTATC-3⬘ Antisense 5⬘-GGTCTGGTCCATCTTGGACAA-3⬘

101

gi: 6981147

Rat Ob-Rb

Sense 5⬘-GCAGCTATGGTCTCACTTCTTTTG-3⬘ Antisense 5⬘-GGTTCCCTGGGTGCTCTGA-3⬘

114

gi: 1526441

Rat Ob-Ra

Sense 5⬘-GTTCCTGGGCACAAGGACTTAAT-3⬘ Antisense 5⬘-ACTGTTGGGAGGTTGGTAGATTG-3⬘

101

gi: 10764822

Mouse LHR

Sense 5⬘-TCTGAAATACTGATCCAGAACACCAA-3⬘ Antisense 5⬘-GGTCCGGATGCCTGTGTTAG-3⬘

111

gi: 7305232

5008

Endocrinology, November 2003, 144(11):5006 –5013


the PCR process. Products were also run on a 2% agarose gel to check for single, correctly sized products.

Housekeeping gene validation To ensure that ␤-actin was an acceptable housekeeping or control gene for this study, it was necessary to determine whether its mRNA levels remained constant across the treatment groups. ␤-Actin mRNA was measured using the ABI GeneAmp 5700 machine and then was normalized to total RNA measurements for each sample. For ␤-actin to be an acceptable control gene, the critical threshold (CT) value per microgram of RNA should not vary statistically across treatment groups. It was also necessary to check that the primers for the housekeeping gene ␤-actin and the other genes of interest had similar amplification efficiencies. Therefore, each primer set was run with serially diluted cDNA, and the resultant CT values were plotted against log dilutions. The slopes of each graph were determined and statistically compared.

Immunohistochemistry Ovaries were collected at 2, 9, 16, and 24 h post-hCG administration, frozen in Jung Tissue Freezing Medium (Leica Instruments GmbH, Nussloch, Germany), and stored at ⫺80 C. Six-micrometer serial tissue sections were cut using a ⫺20 C cryostat (Leica Instruments GmbH). Immunohistochemistry was then performed as previously described (27) using antiserum against rat Ob-Rb [provided by Takashi Murakami; prepared and reported as previously described (30)] diluted 1:2500 in PBS (Life Technologies, Inc.) and 10% normal goat serum (PBS-NGS) as the primary antibody, and biotinylated goat antiguinea pig IgG (Vector Laboratories, Inc., Burlingame, CA) diluted 1:100 in PBS-NGS as the secondary antibody. The sections were then counterstained with hematoxylin, dehydrated, and mounted in DPX (both from BDH Laboratory Supplies, Poole, UK). To ensure antigenic specificity, two types of controls were performed: 1) no primary antibody, replaced with PBS-NGS; and 2) preincubation of antibody with peptide raised against the primary antibody (30) to compete for binding sites. Three sections were examined for each ovary, with both ovaries from six rats per time point.

ANOVA criteria and were then subjected to one-way ANOVA with Tukey’s post hoc analysis. Steroid assay results were also assessed using one-way ANOVA with Tukey’s post hoc analysis. In all cases differences were considered significant at P ⬍ 0.05. All statistical evaluation was performed using the software package SigmaStat for Windows version 2.03 (Jandel Corp., San Ramon, CA).

Results Changes in ovarian weight, RNA content, and housekeeping gene expression

The weights of ovarian tissues were measured immediately before freezing, and total ovarian RNA was measured for each sample collected across the gonadotropin-stimulated cycle (Table 2). The amount of RNA per milligram of tissue increased in response to eCG priming at 26 h post-eCG compared with 4 h post-eCG and decreased immediately before ovulation (9 h post-hCG) compared with 26 and 47 h post-eCG (Table 2). To evaluate ovarian fluctuations in ␤-actin mRNA content across the cycle, expression levels were compared with the total amount of RNA isolated from each ovary. The amount of ␤-actin mRNA per microgram of RNA remained unchanged across the gonadotropin-primed cycle (Fig. 1), indicating that it is constitutively expressed and is therefore an appropriate housekeeping gene and that it is suitable for normalizing data obtained from other genes of interest. The slopes of the plots of CT values vs. log cDNA dilutions were also similar for each primer set (data not shown), indicating that for the data range studied, amplification reactions occurred with the same efficiency. This is further confirmation of the precision of the quantitative RT-PCR assay.

Steroid assays Steroid levels in plasma and culture medium were assayed using a chemiluminescent immunoassay system (Johnson & Johnson Vitros ECI, Orthoclinical Diagnostics, Amersham Pharmacia Biotech, Freiburg, Germany) specific for estradiol and progesterone. The sensitivities of the assays were 10 and 0.3 pmol/liter for estradiol and progesterone, respectively. Rat plasma samples were also assayed using a rat leptin RIA according to the manufacturer’s instructions (Linco Research, Inc., St. Charles, MO), and the sensitivity for this test was 0.5 ng/ml. All assays have inter- and intraassay coefficients of variation less than 8%.

Data analysis To evaluate the effect of treatment group on primer amplification efficiencies compared with the housekeeping gene, t tests were performed on the slopes produced by each primer set. For all genes of interest, mRNA content was calculated for each sample relative to the housekeeping gene, ␤-actin. This was performed using the equation ⌬⌬CT, where ⌬CT is the difference between the gene of interest and the housekeeping gene. Data were further normalized to values from the control group (e.g. immature rats). Each gene of interest is therefore described as the fold change from the control group. To evaluate differences between groups, data underwent log transformation to satisfy

Leptin and leptin receptor expression in gonadotropinprimed whole ovaries

The effect of cycle stage on the abundance of Ob, Ob-Ra, Ob-Rb, and LHR mRNA was evaluated by quantitative RTPCR. All samples were run in triplicate with the five experimental primer sets, and data were normalized to ␤-actin mRNA content. RT-PCR products were run on an agarose gel to check for single, correctly sized products (Fig. 2A). Results are expressed as fold changes from mRNA levels in tissue from immature rats (Fig. 2B). mRNA for all transcripts was found at all time points across the gonadotropin-primed cycle. LHR mRNA levels increased steadily across the priming regimen until 9 h post-hCG, when they declined and then remained low up to 24 h post-hCG. Leptin mRNA expression levels reached their lowest 47 h post-eCG, immediately before hCG administration, and peaked at 2 h post-hCG. The levels then fell to those seen in the immature ovary. Both the

TABLE 2. Weight, total RNA, and RNA per milligram of tissue of whole ovaries Immature

Weight (mg) Total RNA (␮g) Total RNA/mg tissue

10.4 ⫾ 0.2a 19.88 ⫾ 1.2a 1.91 ⫾ 0.11

Post-eCG

Post-hCG

4h

26 h

47 h

2h

9h

24 h

14.02 ⫾ 1.0a,b 24.16 ⫾ 1.8a 1.72 ⫾ 0.07a

19.28 ⫾ 1.2b,c 45.8 ⫾ 4.2b 2.36 ⫾ 0.11a,b

25.88 ⫾ 1.5c 57.25 ⫾ 7.4b,c 2.20 ⫾ 0.25c

24.88 ⫾ 2.8c 54.2 ⫾ 4.7b,c 2.20 ⫾ 0.12d

49.98 ⫾ 5.3d 65.08 ⫾ 2.1c 1.33 ⫾ 0.10b,c,d

53.69 ⫾ 5.5d 95.23 ⫾ 5.4 1.85 ⫾ 0.24

Groups with same letters are not significantly different for weight and total RNA; groups with same letters indicate significant difference for total RNA per milligram of tissue (P ⬍ 0.05). The results represent the mean ⫾ SEM of the average of five or six ovarian pairs.


long functional form of the leptin receptor (Ob-Rb) and the short form (Ob-Ra) increased in their number of transcripts at 9 h post-hCG (⬃7- and 8-fold increases from immature levels; P ⬍ 0.001). After ovulation had occurred (24 h posthCG), Ob-Rb mRNA levels had fallen to their original levels, whereas Ob-Ra mRNA levels remained slightly elevated.

FIG. 1. Levels of ovarian ␤-actin across the gonadotropin-priming regimen, expressed as arbitrary units per milligram of total RNA per ovary (n ⫽ 5– 6 animals/group). Levels of ␤-actin per milligram of total RNA do not change across the cycle (P ⬎ 0.05), indicating that ␤-actin is constitutively expressed and is therefore a suitable housekeeping (control) gene.


Leptin and leptin receptor expression in cultured preovulatory follicles

Dramatic changes in leptin and leptin receptor mRNA were observed in response to hCG. Therefore, to better delineate changes, preovulatory follicles were isolated from eCG-treated rats either before (0 h) or 9 h post-hCG treatment (9 h). Additionally, some isolated follicles from the 0 h group were cultured in vitro with hCG. Treatment with hCG was deemed to be successful if an increase in follicle progesterone production was observed compared with that in untreated follicles. This only occurred at the highest dose of hCG (1000 mIU/ml; P ⬍ 0.05; data not shown), and therefore, these are the only mRNA results shown. Follicles removed from in vivo treated animals either before (0 h) or 9 h after hCG (9 h) displayed LHR, Ob-Rb, and Ob-Ra mRNA production at both stages (Fig. 3A); however, only trace amounts of leptin mRNA were measured, and this was not enough to be quantitated effectively. LHR levels dropped after hCG administration; however, the levels of leptin receptor mRNA remained unchanged. After 9-h culture with 1000 mIU/ml hCG, there was no change observed in LHR, Ob-Rb, or Ob-Ra (Fig. 3B), although there was a tendency for leptin receptor levels to increase in

FIG. 2. A, Single, correctly sized bands were visualized for each set of primers for a representative sample from each time point. M, Molecular weight marker; NTC, no template control. B, Fold differences in mRNA expression of leptin, Ob-Rb, Ob-Ra, and LHR from whole ovaries across the gonadotropin-primed cycle (n ⫽ 5– 6 animals/group). Bars labeled with different letters indicate a statistically different result (P ⬍ 0.05).

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FIG. 3. Fold differences in mRNA expression of LHR, Ob-Rb, and Ob-Ra from preovulatory follicles extracted pre-hCG and 9 h posthCG injection in vivo (A) and preovulatory follicles cultured in vitro for 9 h with 0 or 1000 mIU/ml hCG (B; n ⫽ 5 replicates/group). *, LHR 9 h post-hCG in vivo is statistically different from LHR levels pre-hCG (P ⬍ 0.05).

response to hCG treatment. Again in these follicles, the level of leptin mRNA was too low to be quantitated, although some was detected. Leptin receptor (long form) protein expression

Immunohistochemistry for Ob-Rb showed that this protein is expressed in all compartments of the ovary at all time points studied (Fig. 4). The most intense staining was seen in the endothelial cells (Fig. 4, A and B), and also in the germinal vesicles of the oocytes and then the oocytes cytoplasm (Fig. 4C), The thecal cells, granulosa cells (Fig. 4D), and corpus luteum (not shown) also expressed the leptin receptor protein. The staining appeared less intense in granulosa and thecal cells of smaller follicles than in larger (antral to preovulatory) follicles, although this was not quantitated. Estradiol, progesterone, and leptin levels

Progesterone and estradiol were measured in plasma samples across the cycle (data not shown). Progesterone was found to increase across the cycle, with peak levels measured at 9 h post-hCG. Estradiol levels were highest before hCG administration and then decreased. In follicle culture medium after 9-h treatment with 0 –1000 mIU/ml hCG, progesterone increased only in response to 1000 mIU/ml hCG, and estradiol remained unchanged (data not shown). Leptin was measured in the plasma of rats at various time points across the gonadotropin-priming regimen (Table 3) and displayed low levels across the cycle, with increases observed at 47 h post-eCG and 16 h post-hCG. Discussion

Leptin has been shown to be important in the control of satiety and in maintaining normal reproductive function (5, 31, 32), and the leptin receptor is found on many cells within the ovary, including theca, granulosa, and oocytes (17–19).


Leptin targets these ovarian cells to affect steroidogenesis as well as STAT3 tyrosine phosphorylation and, hence, signal transduction (7, 9, 33, 34). We have previously shown that leptin affects ovulation in vivo and in vitro as well as oocyte maturation, indicating that this hormone has broad-ranging effects within the ovarian environment (27, 28). Leptin mRNA is not detected by sensitive RT-PCR technologies in either human or murine oocytes, yet the protein is clearly present, indicating that it must be produced elsewhere and then transported into the oocyte. Its production by surrounding ovarian somatic cells would present the oocytes with a source of leptin, and many of these cells have been shown to produce leptin (4). Using quantitative RTPCR, we have shown that total ovarian leptin mRNA production increases dramatically 2 h after hCG administration and then declines to unstimulated levels 7 h later. Transcripts for both the long form (Ob-Rb) and one of the short forms (Ob-Ra) of the leptin receptor increase 9 h post-hCG, and tend to decrease after ovulation. We initially expected that the increase observed before ovulation would occur in preovulatory follicles, possibly in the thecal cell layer. What we found, however, was that follicles isolated 9 h post-hCG administration showed no increase in receptor mRNA compared with follicles isolated before hCG, although they did display an expected decrease in LHR mRNA production. We also found that culturing isolated preovulatory follicles with hCG did not affect the mRNA production of either leptin receptors or LHRs. This suggests that the increase in receptor mRNA observed in whole ovaries cannot be accounted for by increases in preovulatory follicular cell production. Leptin mRNA was practically undetectable in preovulatory follicles, implying that its production in the ovary generally occurs in other cell types also. We localized the presence of Ob-Rb protein immunohistochemically and found that there were high levels of receptor protein expression in the endothelial cells of blood vessels within the ovary as well as in the germinal vesicle and cytoplasm of the oocyte and varying levels in the granulosa and thecal cell layers, with intensities in these compartments appearing to increase as the follicles matured. This apparent increase in Ob-Rb protein in larger follicles corresponded to the observation that the mRNA levels increased immediately before ovulation. Protein was also observed within the corpora lutea of recently ovulated ovaries and may be due to production of the receptor by luteinizing granulosa cells. Plasma leptin levels varied across the primed cycle, with significant increases occurring at 47 h post-eCG and 16 h post-hCG, and decreases occurring at 4 h post-eCG and 9 h post-hCG. This contrasts with other studies that have shown either no change in leptin levels (35) or a decrease only before ovulation (36). High levels of leptin receptor mRNA do not necessarily indicate that there will be amplified receptivity to the increased leptin observed immediately before ovulation, although this is a likely scenario. The increase in leptin and leptin receptor expression may be required for oocyte maturation, steroidogenesis alteration, macrophage functionality, etc., at ovulation. An increase in Ob-Ra may be necessary for leptin transport, possibly into the oocyte, as it is known that the oocyte cannot produce its own leptin, but that it has



FIG. 4. Immunohistochemical localization of Ob-Rb in whole rat ovary sections. Ob-Rb staining is found in the endothelial cells of a capillary 24 h post-hCG (magnification, ⫻20). B, Thecal and capillary staining in the ovary 9 h post-hCG (magnification, ⫻40). C, Dark staining is observed in the germinal vesicle of an oocyte in a preantral follicle (magnification, ⫻20). D, Dark staining is observed in the thecal cell layer 16 h post-hCG (magnification, ⫻40). Controls, where primary antibody was preabsorbed with a peptide fragment: E, corpus luteum (magnification, ⫻20); F, preantral follicle (magnification, ⫻20). e, Endothelial cells; t, theca; gv, germinal vesicle; o, oocyte; g, granulosa cells; cl, corpus luteum. TABLE 3. Plasma leptin concentrations (nanograms per milliliter) across the rat gonadotrophin-primed cycle Post-eCG

Immature

0.86 ⫾ 0.28

4h a

0.27 ⫾ 0.27

Post-hCG

26 h b

0.5 ⫾ 0.29

47 h c

1.41 ⫾ 0.34

2h d

0.74 ⫾ 0.23

9h c,f

0.3 ⫾ 0.16

16 h

Groups with the same letter are statistically different (P ⬍ 0.05). The results represent the mean ⫾ each time point.

the ability to produce both Ob-Ra and Ob-Rb mRNA (37), and it has been suggested that Ob-Ra may be involved in transmembrane transportation (25). Leptin receptor levels do not increase in follicles cultured in vitro for 9 h, and this coupled with the fact that preovulatory follicles from in vivoderived ovaries 9 h post-hCG exposure do not exhibit an

1.33 ⫾ 0.18

e

SEM

24 h f

0.38 ⫾ 0.17g

of five or six different animals at

increase in receptor levels either suggest that the increase observed in the whole ovary is not due to an increase in preovulatory follicles, but may be elsewhere in the ovary (for example, where we have displayed staining in theca of other follicles, endothelial cells, and stroma). The decrease in LHR mRNA after hCG administration is

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consistent with findings that LHR levels are regulated in a complex manner, decreasing in response to hCG treatment at 6 h post-hCG, with the lowest levels at 12 h post-hCG (38, 39). It has been suggested that the changes in LHR mRNA expression across the cycle are mainly due to changes in granulosa cell expression of this gene (38). Results from immunohistochemical staining indicate that the leptin produced in the ovary may be acting through functional receptors in all parts of the ovary, specifically at blood vessels and in the oocyte. Leptin may play a role in angiogenesis, as leptin receptors are expressed on vascular endothelial cells (40, 41), possibly by playing a role in matrix remodeling by regulating the expression of matrix metalloproteinases and tissue inhibitors of matrix metalloproteinases (42). Staining observed within the oocytes along with our previous work (27) suggest that leptin may also affect oocyte maturation around the time of ovulation. Staining observed in the thecal cell layer and also in the endothelial cell layers of blood vessels may also be due to macrophage expression of leptin receptors. Macrophages are recruited into these compartments of the ovary in large numbers at the time of ovulation, and it is known that these cells express leptin receptors (13). Leptin affects macrophage functions such as cytokine production (13) and phagocytosis (16), and it is well documented that there are high levels of macrophages in the ovary at the time of ovulation (11). If macrophages are responsible for the change in receptor expression, then this would also explain the lack of increase in receptor mRNA in the cultured follicles, as macrophages are not able to be recruited to these in vitro follicles. It has also been suggested that leptin acts in an autocrine manner to regulate its own expression through the leptin receptor (43). In db/db mice and fa/fa rats (both deficient in functional leptin receptors), there is a high level of circulating leptin that may be attributable to an absence of receptormediated feedback. We suggest that leptin may also act to control the production of its own receptors via actions through existing receptors, as we observed an increase in receptor mRNA production immediately after an increase in leptin mRNA production. There is currently speculation about the role of leptin within the ovary, without any clear evidence to indicate its precise function in this organ. We suggest that leptin has actions on the ovary via its specific receptors at ovulation and/or immediately after ovulation and may act on 1) the oocyte, to promote maturation in readiness for fertilization postovulation; 2) ovarian macrophages or infiltrating monocytes, to promote cytokine production and phagocytosis that occurs during tissue remodeling throughout ovulation and corpus luteum production; and 3) blood vessels within the ovary, to promote angiogenesis during development of the corpus luteum. Our study provides evidence that the production of leptin and its receptors is regulated within the ovary, and we suggest that the leptin produced in the ovary may regulate its own receptor. Acknowledgments We are grateful to Dr. Rebecca Robker, Dr. Robert Gilchrist, and Prof. David Armstrong for their valuable assistance in the preparation of this


manuscript, and to Lesley Ritter and Melinda Jasper for their important technical assistance regarding the real-time RT-PCR work. Received May 22, 2003. Accepted August 6, 2003. Address all correspondence and requests for reprints to: Dr. Robert J. Norman, Reproductive Medicine Unit, Department of Obstetrics and Gynecology, University of Adelaide, First Floor Maternity Building, Queen Elizabeth Hospital, Woodville Road, Woodville, South Australia 5011, Australia. E-mail: [email protected]. This work was supported by a project grant from the National Health and Medical Research Council of Australia.

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