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Reproductive Medicine Unit (P.S.D., K.H.V.d.H., C.R.M., N.K.R., D.T.A., R.J.N.), Department of Obstetrics & Gynecology, The University of Adelaide, The Queen Elizabeth Hospital, Woodville 5011, S.A. Australia; and Department of Obstetrics and Gynecology (D.A.M.), Cedars-Sinai Burns & Allen Research Institute, Los Angeles, California 90048
Address all correspondence and requests for reprints to: Robert J. Norman, M.D., University of Adelaide, Queen Elizabeth Hospital, Department of Obstetrics/Gynecology, Woodville 5011, S.A. Australia. E-mail: robert.norman{at}adelaide.edu.au
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Introduction |
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Leptin messenger RNA and protein are synthesized and secreted from adipose tissue. While some authors indicate the presence of leptin messenger RNA in the ovary (2), others have not confirmed this (3, 4). Several lines of evidence suggest that leptin may be functional in reproductive tissues. Firstly, the leptin protein is found in follicular fluid with levels corresponding to those found in serum (2, 5). Secondly leptin levels in peripheral blood samples vary throughout the menstrual cycle (6, 7, 8), with leptin levels peaking in the luteal phase in a similar manner to 17ß-estradiol and progesterone (6, 9). Thirdly, the expression of the functional leptin receptor in ovarian follicular cells, including human granulosa, theca, and interstitial cells (2, 4, 10), supports the possible direct involvement of leptin in ovarian function. Finally, in vitro studies demonstrate leptin inhibition of insulin-like growth factor I-mediated enhancement of FSH-stimulated estradiol synthesis by rat and human granulosa cells (11, 12) and of LH-stimulated androgen synthesis by bovine theca cells (11).
The present study evaluates the effect of high systemic leptin on the ovulation process. The purpose of this study was to examine the effect of leptin on the ovulatory process, both in vivo, using immature gonadotropin-primed animals, and in vitro, using the perfused rat ovary model. We also set out to establish if the levels of the steroid hormones, progesterone and estradiol, in either model were affected by leptin treatment.
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Materials and Methods |
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Animals
Immature female Sprague Dawley rats aged 21-28 days (from the
University of Adelaide colony) weighing 60–80 g were maintained under
controlled conditions of light (14-h light, 10-h dark), temperature and
humidity, with free access to pelleted food and water. Animals were
handled in accordance with the Australian code of practice for the care
and use of animals for scientific purposes and the animal ethics
committees of both The Queen Elizabeth Hospital and The University of
Adelaide approved all experiments.
In vivo studies
Gonadotropin priming. For the in vivo studies,
follicle growth was induced in immature rats by infusion of hFSH (8 IU)
or saline via Alzet mini-osmotic pumps (Alza Corp., Palo
Alto, CA) releasing 0.5 µl/h. These were implanted sc under
anesthesia on experimental day 1 (13, 14) induced with an inspired gas
mixture of 3% halothane in nitrous oxide and oxygen, followed by 1%
halothane in nitrous oxide and oxygen for anaesthetic maintenance. The
skin was closed with sterile autoclip surgical wound clips
(Becton Dickinson and Co., Franklin Lakes, NJ), and
the rats were placed under a heat lamp until normal activity resumed.
The FSH dose used was based on preliminary dose-response studies in
which concentrations of 4, 8, and 12 IU FSH were used to stimulate
follicle growth. FSH (8 IU) administered over the course of the
experiment produced a mild superovulatory response with ovulation rates
only slightly greater than those of naturally cyclic adult rats of this
strain (13, 14).
Experimental groups. On experimental day 3, at 0800 h, rats from both unstimulated and stimulated groups were injected ip with either leptin (30 µg in 200 µl) or 200 µl saline only, and thereafter at 3 hourly intervals for 15 h. This gave rise to a total of 5 groups of animals: 1) FSH/saline—in which animals received an 8 IU FSH minipump and saline injections on day 3; 2) FSH/leptin—in which animals received an 8 IU FSH minipump and leptin injections on day 3; 3) saline/saline—in which animals received a saline minipump and saline injections on day 3; 4) saline/leptin—in which animals received a saline minipump and leptin injections on day 3; and 5) pair-fed—in which animals were treated with FSH/saline as in group 1 but were fed the same amount of food as that consumed by the FSH/leptin treated animals (group 2). At 1200 h on day 3, 10 IU hCG was administered ip to all groups to induce ovulation. In all groups, body weight and food consumption were measured daily. The dose of leptin administered was based on studies by others (15, 16, 17, 18) that used similar concentrations of leptin. The final dose used over the course of the day was determined using pharmodynamic observations to give an expected concentration of leptin in the high physiological range.
Oocyte and tissue collection. On the morning of day 4, rats were killed using ketamine/xylazine anesthesia and cervical dislocation. The ampulla region was isolated from the ovarian tissue, and the ovulated oocytes were removed and counted from both left and right ovaries. Both ovaries were then dissected free of the ovarian fat and bursa, weighed, and frozen in tissue freezing media using isopentane and liquid nitrogen and stored at -80 C. To determine if leptin affected follicle growth to the preovulatory size, animals stimulated with FSH were killed before the final injection of leptin or saline at 2300 h on day 3. Ovaries from these animals were removed and frozen as described above. In addition, a blood sample was collected by cardiac puncture for plasma leptin, progesterone and estradiol analyses.
Ovarian morphology. Four pairs of ovaries (five for the
pair-fed group) were selected for ovarian morphological analyses on the
basis of mean representative ovulations for each treatment group. Each
pair of ovaries was serially sectioned (6 µm thick). Representative
sections collected every 130 µm were stained with hematoxylin and
eosin, and the numbers of follicles in each of the five antral follicle
classes were counted, as described previously (19). Briefly, antral
follicles were classified on the basis of mean diameters and placed
into one of the five following classes: 275–350 µm (class 1);
351–400 µm (class 2); 401–450 µm (class 3); 451–575 µm (class
4); or 
576 µm (class 5). Class 5 follicles are preovulatory
follicles and are destined to ovulate. Stained sections were visualized
and measured using Video Image Analysis software from Leading Edge Pty.
Ltd. (Adelaide, Australia).
Plasma leptin levels. Leptin levels in plasma samples collected from animals immediately before ovulation were analyzed using one human leptin ELISA kit from DSL Scientific (Webster, TX). This assay does not cross-react with rat leptin. The minimum detection limit of this assay is 0.05 ng/ml and the intraassay CV is 2.9%.
In vitro studies
Gonadotropin priming. For in vitro studies, all
rats received a sc injection of eCG (20 IU) at 1200h on experimental
day 1 to promote the growth and maturation of a first generation of
antral follicles (14).
Surgical isolation of ovaries. On the morning of day 3, following eCG priming, animals were anaesthetized with ketamine/xylazine (67/14 mg/kg BW), and the right ovary was isolated using surgical procedures described in detail previously (20, 21).
In vitro perfusion technique. The ovary was placed in a 30-ml recirculating system, filled with perfusion medium M199 (supplemented with 4% BSA, 50 mg/ml gentamycin, 0.026 M sodium bicarbonate and 0.021 U/ml insulin). Pressure was maintained at 80 mmHg resulting in an average flow of 1.25 ml of media per minute through perfused ovaries. Following connection, ovaries were perfused for 1 h to allow metabolic stabilization of the tissue before the treatments of LH alone (0.1 µg/ml) or LH with leptin (1 µg/ml) were added to the perfusion media. Samples of the circulating medium were taken at 0, 1, 2, 3, 4, 8, and 22 h following commencement of treatments and stored at -20 C for subsequent steroid hormone analyses. The perfused ovaries were removed from the perfusion system at 22 h post treatment, and the surface rinsed with saline to dislodge any adhering oocytes before the number of oocytes found in the perfusion chamber were counted. Two criteria were used to ensure that only properly perfused ovaries were included in the results: firstly, progesterone levels in the first 3 h of the perfusion met specific criteria (22) and secondly, ovaries were examined post perfusion with hematoxylin to determine if vasculature of the ovary was intact.
Progesterone and estradiol assays
Progesterone and estradiol levels from rat plasma and perfusion
media samples were analyzed using a Johnson & Johnson
Vitros ECI Chemiluminescent immunoassay system (Orthoclinical
Diagnostics, Amersham Pharmacia Biotech, Freiburg,
Germany) with sensitivities of 0.3 pmol/liter and 10 pmol/liter,
respectively. The progesterone and estradiol assays have interassay CVs
of <8% and <6%, respectively. Both assays were validated
for rat plasma using a hexane/ethyl acetate extraction.
Statistical analyses
The statistical analyses for the in vivo component of
the study (n 12, unless stated otherwise) were performed using
a parametric ANOVA with Tukey-Kramer multiple comparisons test.
In vitro ovulation data (n 6) were analyzed using an
unpaired two-sided Mann-Whitney test. In vitro progesterone
and estradiol results (n 6) were analyzed using a one-way
ANOVA. In all studies, statistical significance was accepted if
P < 0.05.
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Results |
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). Total
number of oocytes ovulated from left and right ovaries were examined
from the ampullae of rats at 22 h post hCG. Counting of oocytes
revealed that animals primed with FSH/saline ovulated the highest
number of oocytes (15.9 ± 2.0), and animals that did not receive
any FSH (saline/saline) ovulated the least number of oocytes (3.7
± 1.0). The number of oocytes ovulated by pair-fed animals (14.5
± 1.7) was not significantly different from those of the FSH/saline
treated group, whereas leptin treatment of FSH-stimulated animals
caused a significant reduction in ovulation (5.3 ± 1.6)
(P < 0.001). The number of oocytes ovulated following
saline/leptin treatment (2.3 ± 0.9) were not significantly
different to saline/saline treated animals. Both of these groups
ovulated significantly fewer oocytes than animals receiving either
FSH/saline or pair-fed treatments (P < 0.001). Oocytes
ovulated from animals receiving FSH/leptin and those receiving
saline/saline or saline/leptin were not significantly different.
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). Food
consumption and body weight were measured at 0900 h daily. All
groups gained weight before the treatment given on day 3. The day after
leptin or saline ip injections, leptin-treated animals consumed less
food than control animals, and as a result, lost weight between days 3
and 4. Body weight of pair-fed animals was not significantly different
from those of the FSH/leptin-treated group. Ovarian weights of the
FSH-treated animals were significantly different from those animals
receiving saline treatment (P < 0.05); however, leptin
treatment on its own did not affect ovarian weights.
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Ovarian morphology (Fig. 2).
Histological analysis of control and leptin-treated ovaries immediately
before ovulation (Fig. 2A) and after ovulation (Fig. 2B) revealed that
leptin treatment did not alter the numbers of follicles recruited into
the preovulatory stage. Similarly, all other follicle classes (classes
1–4) were unaffected by leptin treatment. A difficulty in
distinguishing between very early corpora lutea and preovulatory
follicles in the postovulation study meant that class 5 follicles
included all follicles that exceed 576 µm. Therefore, both ruptured
follicles that had released oocytes and preovulatory follicles that did
not ovulate are included in the class 5 category of the postovulation
study.
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In vitro studies
Ovulation rate (Fig. 3). The
reduction in ovulation rate observed in vivo was reflected
in vitro. Ovaries perfused with both LH and leptin ovulated
significantly fewer oocytes (1.3 ± 0.6 per ovary) than ovaries
perfused with LH alone (5.7 ± 1.6) (P =
0.014).
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). Both
groups of perfused ovaries showed an immediate increase in the level of
circulating progesterone in response to LH addition (Fig. 4A). The
levels of progesterone then declined after this initial peak, as
previously described (22). No significant difference was seen in
circulating progesterone levels between treatment groups at any time
during the perfusion period. LH addition also resulted in the
stimulation of estradiol to a maximal level within 8 h (Fig. 4B),
after which, levels plateaued for the remainder of the perfusion
period. No significant difference between estradiol levels of the two
treatment groups was observed.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The expression of the long form of the leptin receptor has been detected in granulosa, theca, interstitial, and cumulus oophorus cells (2, 3, 4, 5) of the ovary. The use of the in vitro perfusion system clearly demonstrates that leptin as a cytokine exerts a specific effect on the ovulation process. Leptin injected either systemically in vivo or into the perfusate led to a significant reduction in ovulation rates. In the in vivo study, we used immature animals primed with gonadotropins. This design allowed us to reduce or eliminate the effects of leptin on the hypothalamic-pituitary axis (23) by over-riding endogenous levels of gonadotropins with injected FSH. Leptin was able to inhibit ovulation in these gonadotropin-stimulated ovaries, suggesting that its primary action was on the ovary and not on organ systems outside the hypothalamic/pituitary/ovarian axes. Given that leptin treatment causes a decline in body weight and food consumption (24), a set of pair-fed animals were introduced to compensate for the effects of weight loss and altered acute nutrition. The use of this group clearly demonstrates that leptin directly, as opposed to indirectly through a decrease in body weight, is responsible for the reduction in ovulation seen in these experiments. Ovarian weight was not affected by leptin treatment or by pair feeding, while, as expected, animals treated with gonadotropins had significantly higher ovarian weights when compared with saline-treated controls. Concentrations of leptin measured in the plasma of animals immediately before ovulation were undetectable, in agreement with the short half-life (24.9 ± 4.4 min) of the leptin protein (25, 26). Biological activity was clearly present however, shown by the effect on body weight.
We have extensively characterized the value of the in vitro perfused ovary system with a variety of gonadotropins and cytokines (27, 28). CG priming of animals, followed by surgical removal of their ovaries, selective cannulation, and injection of LH allows for investigation of factors acting directly on the ovary to influence the process of ovulation. The injection of leptin at the same time as LH clearly shows that this protein was able to inhibit the ovulatory process in the isolated ovary, thereby supporting the suggestion that leptin has a direct action on the ovary. In both in vivo and in vitro studies, there were no changes seen in steroid secretion. This finding is consistent with previous research in rat ovarian granulosa cells, when in the absence of growth factor augmentation, leptin is unable to affect steroid production (12, 29). In the presence of high levels of IGF-I, the sensitizing effects of FSH enhance progesterone and estradiol production of rat granulosa cells (11). However, high levels of leptin have been postulated to block the stimulatory effects of IGF-I on rat granulosa cell estradiol, but not progesterone, production without altering the effects of FSH alone (11).
Our results clearly show that leptin can reduce ovulation rates in vivo and in vitro when administered acutely. While systemic leptin in the live animal may potentially affect many systems, the use of an isolated ovary indicates that leptin action is partially, if not totally, on the ovarian tissue. The mechanism of action however, remains uncertain. The decrease in ovulation observed in the study could not be attributed to an apparent decrease in the number of preovulatory follicles as both the number of these follicles and ovarian weights were unchanged in leptin-treated animals. It is possible that oocytes are trapped inside the follicles, but this was not detected with histopathology. If this is the case, then leptin administration may interfere with LH action, thereby preventing the release of the oocytes from the preovulatory follicle. In vitro studies in dispersed ovaries performed using IGF-I suggested that leptin can impair LH action (5).
The processes within the ovary that follow the LH surge and lead to ovulation have been well characterized (30, 31, 32, 33). LH initiates an increase in interleukin-I production as well as the induction of a variety of other cytokines, collagenases, plasminogen activators, and enzymes (33). Leptin could affect any of these processes. We have been particularly interested in the role of leukocytes in the ovulation process, where their importance has been clearly established (34, 35, 36, 37) with numbers in the rat ovary increasing immediately before ovulation (36). Leukocytes can release many factors, such as plasminogen activators, collagenases, proteases and cytokines that may assist in follicle wall degradation for the release of the oocyte (38). Any inhibition of leukocyte infiltration into the ovary or preovulatory follicle could result in the failure of the follicle to ovulate. Leptin has recently been shown to be capable of interactions with the immune axis (39) and may therefore influence the infiltration of macrophages or other leukocytes into the preovulatory follicle, hence disrupting the ovulatory process. Examination of leukocyte numbers in the ovaries of leptin-treated animals is in progress and preliminary results show a decrease in neutrophil infiltration in thecal tissue.
Leptin has prominent effects on the reproductive axis and is able to reverse the sterility observed in the leptin-deficient obese (Lepob/Lepob) mouse (17, 18). It has been shown that serum leptin levels proportionally correlate with body mass index and percentage body fat in women (40, 41). Furthermore, obesity has also been linked to reproductive dysfunction. The leptin levels found in grossly obese women range to 100 ng/ml (41). The leptin levels used in this study could be expected to exceed the physiologically high leptin levels observed in obese women (41, 42). Our data and other reports (10, 12) indicate that elevated leptin concentrations are able to exert a direct inhibitory effect on ovarian function. This research, therefore, provides a new dimension to studies on leptin and reproduction by indicating that acute administration of leptin, both in vivo and in vitro, can impair the ovulatory process. This may have clinical correlates in the observation that overweight women (with high leptin) are more prone to ovulatory disorders that cannot be entirely explained by the hypothalamic-pituitary axis. However, it is important to acknowledge that the ovulatory dysfunction experienced by most obese women may be due to the insensitivity to endogenous leptin (41) and the chronically high levels of leptin may not mirror the effects seen by acute exogenous leptin administration.
In summary, we have investigated the effect of high leptin levels both in vivo and in vitro. In both studies we found significant decreases in the number of oocytes ovulated, without any apparent effect on steroid levels. While leptin treatment decreased appetite and body weight, pair feeding of animals did not alter the results, indicating that leptin alone affects ovulation. Our evidence suggests a direct effect of leptin on the ovary, independent of alteration in the amount of gonadotropins, and other circulating growth factors. Further investigations will concentrate on the effect of physiological levels of leptin on the ovary and the effects of leptin levels on some of the ovulatory mediators and cells that are recruited into the ovary at the time of ovulation.
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Acknowledgments |
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Footnotes |
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2 Supported by a Reproductive Medicine Laboratories Postgraduate
Scholarship, The University of Adelaide.
Received November 30, 1999.
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