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RESEARCH |
1 Dpto de Química Biológica, Facultad de Ciencias Exactas y Naturales (FCEyN), Universidad de Buenos Aires (UBA), Buenos Aires, Argentina and 2 Centro de Estudios Farmacológicos y Botánicos (CEFYBO), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Facultad de Medicina, UBA, Paraguay 2155, 16 Piso, Buenos Aires C1121ABG, Argentina
Correspondence should be addressed to A G Faletti; Email: agfaletti{at}yahoo.com.ar
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Abstract |
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Introduction |
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The effects of leptin on ovulation are contradictory and both stimulatory and inhibitory actions on ovarian function have been described. In addition to the effects on the hypothalamic–pituitary axis, we can mention some negative actions: (i) leptin can directly suppress insulin, insulin-like growth factor-I (IGF-I), transforming growth factor-ß and glucocorticoid-induced steroidogenesis of ovarian granulosa cells of rat (Zachow & Magoffin 1997, Barkan et al. 1999, Brannian et al. 1999, Zachow et al. 1999) or human (Agarwal et al. 1999) and (ii) acute administration of leptin to immature gonadotrophin-primed rats inhibits ovulation (Duggal et al. 2000, 2002). Likewise, leptin is able to produce some positive effects: (i) leptin accelerates the onset of puberty in rodents (Ahima et al. 1997, Almog et al. 2001) and humans (Clément et al. 1998, Strobel et al. 1998); (ii) leptin induces ovulation in GnRH-deficient mice (Barkan et al. 2005) and eCG/hCG-primed rats (Roman et al. 2005).
Nevertheless, the precise mechanism by which leptin affects the ovulatory process is completely unknown. Ovulation is a complex process involving gonadotrophins, steroid hormones and many mediators common to inflammatory reactions, such as cytokines, prostaglandins, leukotrienes, plasminogen, nitric oxide and histamine. Prostaglandins (PGs) and nitric oxide (NO) are of particular interest, as they have been shown to play a role in follicle rupture (Brännström & Janson 1991, Ellman et al. 1993, Shukovski & Tsafriri 1994). In a previous study, we have demonstrated that during the ovulatory process, the increase in ovarian nitric oxide synthase (NOS) activity results in an increase in NO, which stimulates PGs production and enhances the inflammatory process, facilitating follicle rupture (Faletti et al. 1999).
Therefore, the aim of the present study was to investigate the negative action of leptin on some of these ovarian preovulatory factors during the ovulatory process by performing both in vivo studies using prepuberal rats stimulated with gonadotrophins and in vitro studies, using ovarian explants and follicle cultures as biological models. The expression of leptin receptors in the ovarian tissues was also evaluated.
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Drugs and chemicals
hCG, PGE2, recombinant rat leptin, progesterone and proteases inhibitors were purchased from Sigma-Aldrich. eCG was obtained from Syntex SA (Buenos Aires, Argentina). [3H]-PGE2 (181 Ci/mmol) and [1,2,6,7-3H]-progesterone were obtained from Amersham Pharmacia Biotech. The western blotting reagents were obtained from Sigma-Aldrich and Bio-Rad Laboratories.
In vivo studies
Animals were injected i.p. with 10 iu eCG (in 0.10 ml saline) to induce the growth of the first generation of preovulatory follicles. Forty-eight hours later, the animals were injected i.p. with 10 iu hCG (in 0.10 ml saline) to induce ovulation, which usually occurs within 12–14 h after hCG administration in this rat colony.
To study the serum concentration of leptin and the expression of leptin receptors (Ob-R), animals were killed by decapitation at different times during the gonadotrophin treatment. Trunk blood was collected, serum was harvested and stored at –20 °C until RIA studies. Ovaries were dissected out quickly, frozen on dry ice and stored at –70 °C. Eight or ten animals were killed at each time, resulting in 16–20 ovaries/group.
To study the effect of high level of leptin during the ovulatory process, rats received five i.p. injections of either recombinant rat leptin (5 µg/0.15 ml PBS-BSA) or PBS-BSA (control) 1 h before hCG administration and at intervals of 150 min. To study the ovulation rate, some animals (four to five per group) were killed 20 h after the hCG injection by cervical dislocation. The remaining rats (four to five per group) were killed 10 h after the hCG injection by decapitation and trunk blood was collected. Sera were harvested and stored at –20 °C until assayed for progesterone by specific RIA. The ovaries were dissected out quickly, weighed and frozen on dry ice and stored at –70 °C. One ovary from each animal was used to determine prostaglandin E (PGE) content by RIA and the other one to measure the expression of leptin receptors by western blot analysis. Previously, we have shown that gonadotrophin administration increases ovarian NOS activity and PGs content, and that these increases peak at 10 h after hCG administration (Faletti et al. 1999). Therefore, we chose this time as a preovulatory moment when NOS activity and PGs content are at their highest levels before ovulation occurs. The experiments were repeated at least two times.
Ovulation rate
Twenty hours after hCG injection, the rats were killed by cervical dislocation, ovaries were immediately removed and oviducts were dissected and examined by means of a stereoscopic microscope to assess the number of oocytes present within, as described previously (Faletti et al. 1995).
Ovarian PGE content
PGE was extracted from the ovaries as described previously (Faletti et al. 1997). Briefly, one ovary from each animal was homogenised in absolute ethanol, centrifuged at 1000 g and the supernatants evaporated to dryness. The residues were stored at –70 °C and reconstituted before being assayed. PGE was quantified by RIA as in previous studies (Faletti et al. 1997) using rabbit antiserum (P5164) from Sigma-Aldrich. The sensitivity of this assay was 15 pg/ml and the cross-reactivity of PGE2 was 100% with PGE1 and lower than 0.1% with other prostaglandins. The intra- and interassay coefficients of variation were 8.2 and 12% respectively.
Western blot analysis
Soluble tissue extracts were prepared as described previously (Faletti et al. 2003). Briefly, ovaries were homogenised in 20 mM ice-cold Tris–HCl buffer (pH 7.4), containing 0.25 mM sucrose, 1 mM EDTA, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 100 µg/ml phenylmethylsulfonyl fluoride and 10 µg/ml trypsin inhibitors. The homogenates were sonicated and centrifuged at 7800 g at 4 °C for 15 min and protein concentration in the supernatant was determined by the Bradford method with BSA as the standard. Homog enates were boiled for 5 min in buffer containing 0.3% (w/v) bromophenol blue and 1% (v/v) ß-mercapto-ethanol. Equal amounts of protein (100 µg) were loaded onto 4% (w/v) 0.125 M Tris–HCl (pH 6.8) stacking polyacrylamide gel, followed by a 7.5% (w/v) 0.375 M Tris–HCl (pH 8.8) separating polyacrylamide gel. Following electrophoresis, proteins were transferred to PVDF membrane (Bio-Rad Laboratories) for 60 min in a cold chamber using a Bio-Rad transblot apparatus. Membranes were first blocked at 4 °C overnight in Tris–HCl:saline (50 mM Tris–HCl:150 mM NaCl (pH 7.5)) containing 5% (w/v) of milk powder, and then incubated at 4 °C overnight with antibody raised in rabbit against Ob-R (H-300) (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The final dilution of antibody was 1:200. The membranes were washed four times for 15 min each in Tris–HCl:saline containing 0.1% (v/v) Tween-20 (pH 7.5; TTBS). Negative controls were carried out by omitting the incubation with the primary antibody. No bands were detected. Then, the sections were incubated for 1 h at room temperature with goat anti-rabbit IgG (1:2500) as the secondary antibody (Santa Cruz Biotechnology). The antibody was then washed off in TTBS and the immunoreactive bands were visualised using chemiluminescence detection reagents (Sigma-Aldrich) and exposed to Kodak X-OMAT film. Before reuse, membranes were stripped, blocked and reprobed according to the manufacturer’s instructions. Membranes were reprobed with anti-actin antibody (A2066, Sigma-Al-drich). Molecular weight standards (Kaleidoscope St; Bio-Rad Laboratories) were run under the same conditions to identify the protein bands. Blots were scanned using a scanning UMAX Astra 12205 and densitometry was analysed using a Dekmate III Sigma Gel software package (Jandel Scientific software). The data were normalised to ß-actin protein levels in each sample to account for procedural variability.
In vitro studies
Ovarian explant culture
At days 26–28, animals were injected i.p. with 10 iu eCG (in 0.10 ml saline) to induce the growth of a first generation of preovulatory follicles. Forty-eight hours later, the animals were injected i.p. with 10 iu hCG (in 0.10 ml saline) to induce ovulation and were killed by cervical dislocation 4 h later. Both the ovaries were immediately removed and dissected free of fat and bursa, and were cut into pieces of approximately equal size (four slices/ovary). Ovarian slices (four slices/well) were placed in 24-well plates containing Dulbecco’s modified Eagle medium (DMEM)/F12 (1:1; Bio-Rad Laboratories) medium with 25 mM HEPES, 100 U/ml penicillin, 100 µg/ml streptomycin, 0.5 µg/ml fungizone and 2 mM L-glutamine. Ovarian slices were incubated in a final volume of 500 µl/well in either the presence or the absence of leptin (30–500 ng/ml) at 37 °C in a humidified atmosphere (5% CO2:95% O2) for 4 h. The dose of leptin used in these studies was obtained from previous reports (Spicer & Francisco 1997). After the incubation period, ovarian tissues were recovered, weighed and the protein content was determined by the Bradford method with BSA as the standard. Culture media were stored at –20 °C until assay for progesterone, PGE and nitrite (as a stable metabolite of NO production) concentrations. At least three independent experiments were run for each culture condition using different ovarian tissue preparations.
Culture of ovarian follicles
At days 26–28, animals were primed with 10 iu eCG i.p. (in 0.10 ml saline) to induce the growth of a first generation of preovulatory follicles and 48 h later, the animals were killed by cervical dislocation. Preovulatory follicles (> 550 µm in diameter) were dissected from the ovaries with the aid of a stereomicroscope and fine forceps. Approximately 126 preovulatory follicles were obtained from 12 ovaries and these were collected and pooled. Follicles were placed in 24-well containing DMEM/F12 (1:1) medium (Bio-Rad Laboratories) with 25 mM HEPES, 100 U/ml penicillin, 100 µg/ml streptomycin, 0.5 µg/ml fungizone and 2 mM L-glutamine. Five follicles per well were incubated in a final volume of 500 µl fresh medium in either the presence or the absence of leptin (30–500 ng/ml) in combination with follicle-stimulating hormone (FSH) (100 ng/ml) or luteinizing hormone (LH) (100 ng/ml) at 37 °C in a humidified atmosphere (5% CO2:95% O2) for 4 h. Both the doses of leptin (Spicer & Francisco 1997) and gonadotrophins (Carnegie & Tsang 1984, Duggal et al. 2002) used in these studies were obtained from previous studies. After incubation, culture media (without follicles) were stored at –20 °C until assayed for progesterone, PGE and nitrite concentrations. At least three independent experiments were run for each culture using different preparations.
Hormone assays
Serum concentration of leptin was quantified by RIA as described previously (Roman et al. 2005) using a highly specific anti-mouse leptin provided by Dr A F Parlow (NHPP, Terrance, CA, USA). Recombinant rat leptin was used for standards and serial dilutions of the samples showed parallelism with the standard curve. The limit of sensitivity was 50 pg/ml and the intra- and interassay coefficients of variation were 7.5 and 9.5% respectively. Results were expressed as pg/ml serum.
Progesterone was quantified by RIA in both serum samples extracted with diethyl ether and culture medium. The progesterone antiserum was kindly provided by Dr G D Niswender (Colorado State University, Fort Collins, CO, USA). The sensitivity of these assays was 15 pg/ml. The cross-reactivities were < 2.0% for 20
-dihydro-progesterone and deoxy-corticosterone and 1.0% for other steroids normally in the serum. The intra-and interassay coefficients of variation were 7.5 and 10.5% respectively.
Nitrites assay
Levels of nitric oxide metabolites were measured in the media of both ovarian explants and follicle cultures, using the Griess reagents as described previously (Polisseni et al. 2005).
Statistical analysis
All data are expressed as mean ± S.E.M. The difference between two groups was analysed using Student’s t-test. Comparisons between more than two groups were performed using a one-way ANOVA and Student–Newman–Keuls multiple comparison test. Differences between groups were considered significant when P < 0.05.
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). Densitometric analysis revealed that the administration of eCG induced increase of the expression of Ob-R protein in comparison with that obtained from animals without treatment, being significant only at 48 h postinjection (Fig. 2B). After the hCG administration, the expression of leptin receptors increased again to reach a maximum. Twenty-four hours after hCG administration and after ovulation occurred, the expression decreased returning to values similar to those obtained at 0 h after hCG administration.
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. No differences were found in either food intake (data not shown) or body weight between animals treated with leptin and control animals along this period, but ovarian weight was significantly decreased (Table 1). We examined the ovulation rate 20 h after the hCG administration. The number of oocytes within oviducts was significantly lower in rats treated with leptin than that in control animals (20 ± 3 vs 11 ± 2; P < 0.05; Table 1).
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).
In vitro studies
To investigate the source of the inhibition of ovulation by leptin, the concentration of PGE and progesterone were assessed in ovarian explants cultured for 4 h in either the presence or the absence of leptin and expressed as pg per mg wet weight. Since nitric oxide is involved in ovulation in rats as it stimulates the production of PGs (Faletti et al. 1999), NO production in these cultures was also quantified by measuring the concentration of nitrites as stable metabolites of NO and expressed as ng per mg wet weight. Ovarian explants were obtained from immature rats primed with eCG/hCG to closely imitate the conditions of in vivo experiments. The direct effect of recombinant rat leptin on ovarian production of PGE, nitrites and progesterone is shown in Fig. 3. Addition of 300 ng/ml or more of leptin led to a significant inhibition of PGE and nitrites production. Neither PGE nor nitrites concentrations were significantly affected by the presence of lower concentrations of leptin when compared with those of controls (P > 0.05; Fig. 3A and B). All doses of leptin assayed also reduced progesterone production in comparison with that of control (Fig. 3C). When the results were expressed per mg protein, the responses obtained with these factors were the same (data not shown).
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and 5 respectively. As expected, both FSH and LH (100 ng/ml) stimulated PGE, NO and progesterone production (P < 0.05). All doses of leptin assayed significantly reduced the production of both PGE and progesterone induced by FSH or LH, but not that of progesterone production when the concentration of leptin was 100 ng/ml in presence of LH (Fig. 4C). These inhibitory effects were not total in most of the concentrations assayed, because neither PGE nor progesterone production returned to basal values. Leptin was able to abolish completely the FSH-stimulated NO production in all doses probed (Fig. 5B), but it only reduced the LH-stimulated NO production at 100 ng/ml (Fig. 4B).
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shows that acute leptin treatment did not significantly alter the ovarian leptin receptor expression in comparison with that of the buffer control, when a densitometric analysis was performed (Fig. 6B).
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Discussion |
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Duggal et al.(2000) have demonstrated that acute leptin treatment inhibits ovulation. In other study, these authors investigated the cause of this inhibition, but they could not demonstrate what ovulatory factors were involved in this negative action (Duggal et al. 2002). Therefore, the action of leptin on some ovarian preovulatory factors were evaluated during the ovulatory process by performing both in vivo studies with prepuberal rats stimulated with gonadotrophins and in vitro studies, using ovarian explants and follicle cultures as biological models. The purpose of the treatment design was to maintain high levels of leptin after hCG administration to avoid the fall of this protein observed in circulation before ovulation. The administration of five doses of 5 µg leptin, at 150-min intervals, to immature eCG/hCG-primed rats during the ovulatory process, was sufficient to inhibit the number of ovulated oocytes. This result confirmed those previously obtained by Duggal et al.(2000), although they administered higher doses at different intervals. Since the administration of leptin was during the light period (between 0700 and 1800 h), it was not necessary to use a group of rats with pair-fed to the leptin-treated animals, because no differences were found in either food intake or body weight between this group and control animals along this period.
It was of interest to study the action of acute administration on some ovarian mediators involved in the ovulatory process. Prostaglandins and nitric oxide are of particular interest, as they have been shown to play a role in follicle rupture (Brännström & Janson 1991, Ellman et al. 1993, Shukovski & Tsafriri 1994, Faletti et al. 1999). The ovarian concentration of PGs increases after LH surge and gonadotrophin stimulation (Brown & Poyser 1984, Faletti et al. 1995). The present study investigated the effect of high levels of leptin on ovarian PGE production by performing in vivo and in vitro assays. Preovulatory PGE content was significantly inhibited in ovaries from rats primed with eCG/hCG and treated with acute administration of leptin. This result was confirmed with in vitro studies, because the concentration of this prostanoid was reduced by the presence of leptin in the culture media of ovarian explants and FSH- or LH-stimulated preovulatory follicles from rats primed with gonadotrophin. In a previous study, it has been shown that gonadotrophin administration resulted in an increase in nitric oxide synthase activity and this increase results in an increase in NO, which stimulates prostaglandins production and enhances the inflammatory process, facilitating follicle rupture (Faletti et al. 1999). The present study investigated whether high levels of leptin were able to alter directly the ovarian production of nitric oxide, by measuring the concentration of nitrites, as stable metabolites, in the same culture media of ovarian explants and preovulatory follicles where PGE was determined. The production of nitrites was reduced by the presence of leptin in the cultures assayed, although this result was not significant at all the concentrations assayed. These findings appear to contrast with those reported by Huang et al.(2005), who have found that the inhibitory effect of leptin on human granulose cells is mediated by NO, because the exposure of human GCs to leptin at concentrations of 10 ng/ml decrease the IGF-I-stimulated or IGF-I plus FSH-stimulated 17ß-oestradiol production, and the presence of L-NAME (NG-nitro-L-arginine methyl ester), an inhibitor of NOS, in the culture medium significantly attenuates this negative effect. Furthermore, the concentrations used by these authors (3–30 ng/ml) induce a dose- and time-dependent NO production, but they were lower than those used in our studies. Without considering the biological differences between both cellular systems used, it would not be the first time that leptin is able to produce a dual effect on the biological function. Recently, we have found that a chronic treatment with low doses of leptin induces a swift increase in the ovarian endothelium NOS expression in comparison with that of control animals and this treatment completely reverses the inhibitory effect on this protein expression produced by a severe food restriction (Roman et al. 2005).
A direct inhibitory action of leptin on steroid hormone secretion has been demonstrated independently by different groups in the ovary (Spicer & Francisco 1997, 1998, Zachow & Magoffin 1997, Agarwal et al. 1999, Barkan et al. 1999, Zachow et al. 1999, Ghizzoni et al. 2001, Kikuchi et al. 2001) and other tissues (Tena-Sempere et al. 2001, Cameo et al. 2003). The mechanism involved for such inhibitory action has not been completely clarified, but it has been suggested that this effect is mediated by a modulation on transcriptional factors, such as StAR and P450scc (Tena-Sempere et al. 2001) or overexpression on c-Jun (Barkan et al. 1999). In this paper, leptin treatment also induced significant inhibition in serum level of progesterone and again, this result was confirmed by in vitro studies where the concentration of progesterone was reduced by the presence of leptin in cultures of ovarian explants and preovulatory follicles. Again, the findings of both stimulations by low doses and inhibition by high doses of leptin in ovarian cells in vitro concur with observations by Ruiz-Cortés et al.(2003), where the effects of leptin are shown to be biphasic with regard to stimulation and inhibition of progesterone synthesis. These authors have reported that leptin modulates steroidogenesis in a biphasic manner via STAT-3. Some in vivo studies performed with acute high doses have reported that leptin may have inhibitory effects on ovarian factors (Duggal et al. 2000). But other studies have also demonstrated positive action with chronic treatment with leptin, since it has been able to accelerate the onset of puberty in rodents (Ahima et al. 1997, Almog et al. 2001) and human (Garcia-Mayor et al. 1997, Strobel et al. 1998). We have found in a previous study that a chronic administration of low leptin level is able to enhance the ovulatory process. These results are consistent with those obtained by Barkan et al.(2005), who have found that leptin is able to mimic FSH and LH actions or to induce an LH-independent ovulation. All these data suggest that both the dose and timing of leptin administration are critical to obtain either a positive or a negative response.
Functional leptin receptors are expressed in the ovary of numerous species, including humans (Cioffi et al. 1996, Karlsson et al. 1997), mice (Kikuchi et al. 2001) and rats (Zamorano et al. 1997, Zachow et al. 1999). The expression of ovarian leptin receptor was evaluated at different times, after eCG/hCG injection, by western blot analysis to study the effect of the gonadotrophin treatment. This expression varied across the gonadotrophin treatment, with significant increases 48 h after eCG administration and 10 h after hCG administration that tend to reduce after ovulation. These results agree with those obtained by Ryan et al.(2003), who have used quantitative RT-PCR to measure leptin receptor expression. All these data confirm that the production of leptin and its receptors is regulated within the ovary by gonadotrophins, indicating a possible involvement in several ovarian functions such as the ovulatory process. Further studies are necessary to determine which form of this receptor protein is involved and how this expression is regulated.
We initially expected that the increase in the expression of leptin receptor before ovulation would be blocked or partly attenuated by the acute leptin treatment. However, we found that this treatment did not show significant alteration in its expression. This suggests that the inhibitory effect of leptin on the ovulatory process in our in vivo studies was not mediated by changes in the content of receptors protein at ovarian level, at least at the concentrations assayed. Further studies are necessary to clarify this point.
Finally, we firmly believe that leptin may be regulating the ovulatory process by at least in part, modulating prostaglandins, nitric oxide and steroids, since when the levels of leptin are low, there is a positive correlation between leptin and ovarian functions (Roman et al. 2005), but when these levels are high, as in the present study, there is a negative correlation among them.
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Footnotes |
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References |
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P < 0.01 versus –48 h (ANOVA and Student–Newman–Keuls multiple comparison test).
P < 0.001; a. versus LH, b. versus control (ANOVA and Student–Newman–Keuls multiple comparison test).
