| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Eutheria Foundation,3 Cross Plains, Wisconsin 53528
Pathobiological Sciences,4 University of Wisconsin, Madison, Wisconsin 53706
ABSTRACT
The effects of ultrasound morphology, vascularity, and follicular-fluid hormones of the preovulatory follicle on oocyte recovery rate and on follicle and oocyte maturity rates were studied for 60 spontaneous and solitary preovulatory follicles in mares. An ovulation-inducing dose of hCG was given when the follicle was
32 mm (Hour 0), and a procedure for oocyte recovery was done 30 h later (Hour 30). Between Hours 0 and 30, diameter of the follicle increased less and circulating estradiol (E2) concentrations decreased more in groups with successful versus nonsuccessful oocyte recovery and in groups with mature versus immature recovered oocytes, as indicated by significant interactions of group and hour. Significant differences in blood-flow end points between groups were not detected. At Hour 30, the frequency of granulosa serration, an indicator of impending ovulation, was higher (P < 0.001), and the number and expansion of granulosa cells in the lavaging fluid, indicators of follicle maturity, were greater in the oocyte-recovery group and in the oocyte-mature group. Follicular-fluid concentrations of E2, progesterone, and free insulin-like growth factor (IGF) 1 were not different between the oocyte-recovery and -nonrecovery groups. Concentration of progesterone was significantly greater, and E2 and free IGF1 were less in the oocyte-mature than in the immature groups. Results indicated that the post-hCG oocyte-recovery and oocyte-maturity rates were positively affected by follicle maturity. Greater follicular-fluid progesterone and lower E2 and free IGF concentrations were associated temporally with maturation of the oocyte but not with maturation of the follicle.
apoptosis,, blood flow,, follicle,, growth factors,, mares,, oocyte development,, oocytes,, ovarian steroids,, preovulatory follicle,, steroid hormones
Transvaginal aspiration of follicular fluid followed by lavage of the preovulatory follicle is used commonly in mares for oocyte recovery for clinical and experimental purposes [1]. Most mares ovulate when the follicle is 40–45 mm. A common procedure for oocyte recovery involves hCG treatment when the follicle is 33–36 mm and recovery 24–36 h after hCG treatment [2, 3]. Oocytes recovered 24 h after hCG treatment are usually immature and may be cultured for 12 h before transfer to a recipient, whereas oocytes recovered at 36 h are usually mature and can be transferred without culture [1, 2]. Recent studies have demonstrated that in vitro matured equine oocytes can develop efficiently into viable embryos [4]. Nuclear maturation is indicated by extrusion of the polar body at the metaphase II stage, indicating that the oocyte is ready for fertilization [1].
Oocyte recovery is considered difficult in horses, especially when the follicle is immature [1], owing to the cumulus oophorus's broad base of attachment and minimal projection into the antrum [5] and the extension of cumulus-cell processes into a thecal pad [6]. Reported oocyte recovery rates from the preovulatory follicle after hCG treatment have ranged from 65% to 83% [3, 7–10]. Oocyte recovery rates were not significantly different when done at 24 h (66%) and 35 h (73%; [2]) or at 22 h (75%) and 33 h (82%; [3]) after hCG treatment.
Several discrete structural changes that are detectable by B-mode ultrasonography occur in the follicle as ovulation approaches and are indicators of follicle maturity and the imminence of ovulation [11]. Serration of the granulosa becomes distinct, beginning about 6 h before ovulation [11] in the follicle wall opposite the future site of follicle rupture [12]. Other indicators of impending ovulation include a decrease in follicle turgidity and the development of an apical area and echoic spots that float in the antrum [11]. Blood flow increases in the wall of the preovulatory follicle during the 36 h after hCG treatment, based on an increase in the percentage of the circumference of the follicle wall containing color-Doppler signals for blood flow [13]. In a recent study [14], mares were treated with hCG when the preovulatory follicle was 34–37 mm and were bred 30 h later. Percentage of follicle circumference containing blood-flow signals at the time of breeding was greater for mares that became pregnant than for mares that did not. The relationships of changes in follicle vascularity to success in recovering an oocyte and to the maturity of a recovered oocyte have not been reported.
Minimal information is available on follicular-fluid concentrations of steroids in preovulatory follicles of mares or the relationship between steroid concentrations and follicle maturation, oocyte recovery, and oocyte maturation. No differences in intrafollicular estradiol (E2) concentrations were found between mares treated versus not treated with hCG at 35 mm and sampled 28–32 h later, but progesterone (P4) and testosterone were higher in the treated mares [15]. Using a similar hCG-treatment protocol, the proportion of follicles with low E2 concentrations increased in association with nonrecovery of an oocyte [9]. Nuclear and cytoplasmic changes during final maturation of equine oocytes were associated with an increase in P4 in the follicular fluid, whereas E2 remained consistently high [16]. However, in another study with preovulatory follicles (>30 mm), no relationship was found between E2, P4, and testosterone concentrations and oocyte recovery or nuclear maturation [17]. Preovulatory changes in circulating hormones include a decrease in E2 beginning 2 days before ovulation [18], but P4 in hCG-treated mares does not begin to increase in the circulation until approximately 12 h after ovulation [19]. Plasma E2 decreases immediately after an hCG injection given when the follicle is approximately 35 mm [13].
The purpose of the present experiment in mares was to characterize the structural, vascular, and hormonal factors of the preovulatory follicle associated with recovery versus nonrecovery of oocytes 30 h after hCG treatment and maturity versus immaturity of the recovered oocytes.
Animals were handled in accordance with the United States Department of Agriculture Guide for Care and Use of Agricultural Animals in Research. Nonlactating ponies and apparent pony-horse crosses, 3–20 yr of age and 260–480 kg body weight, were used. The study was done during August and September in the Northern Hemisphere. All mares were in their ovulatory season at the time of the procedure for oocyte recovery, as indicated by ovulation during the next cycle. Mares were kept under natural light in an open shelter and outdoor paddock, were maintained on alfalfa/grass hay, and had access to water and trace-mineralized salt. The mares were selected for docile temperament and no apparent abnormalities of the reproductive tract, based on transrectal ultrasound examinations [20]. Mares with a growing 28-mm follicle 15 days after ovulation were scanned daily by B-mode ultrasonography until a 32-mm follicle was detected. Estrous cycles were not used if the 32-mm follicle showed ultrasonographic signs of impending ovulation [11], the corpus luteum was large with an echotexture characteristic of a functional structure [20], two dominant follicles (>28 mm) were present, the score for endometrial edema was characteristic of diestrus (score < 3; [21]), or if ovulation occurred before the oocyte-recovery procedure 30 h later. After removal of estrous cycles, 60 cycles in 37 mares remained. Each selected mare with a 32-mm follicle was given an i.v. injection of 2500 IU of hCG (Hour 0 [Chorulon, Intervet International, Boxmeer, The Netherlands]), and oocyte recovery was attempted 30 h later (Hour 30). Follicular data and blood samples for the E2 assay were collected just before the hCG injection and just before the oocyte-recovery procedure.
To generate optimal ultrasound images, mares were sedated during transrectal scanning with a minimal dose of detomidine hydrochloride (1 mg/mare, i.v.; Dormosedan, Pfizer Animal Health, Philadelphia, PA). A duplex B-mode and pulsed-wave color-Doppler ultrasound instrument (Aloka SSD-3500; Aloka, Wallingford, CT) equipped with a finger-mounted 7.5-MHz convex-array transducer (UST-995–7.5) was used. For B-mode, the brightness and contrast controls of the monitor and the gain controls of the scanner were standardized to constant settings [22]. The color mode was used to display signals of blood flow in vessels of the follicle wall as described [13, 23]. For maximum detection of blood flow by color signals without aliasing, the velocity range was set at 10 cm/s. The color-gain setting was kept constant. The B-mode and color-Doppler end points were evaluated while the entire follicle was being scanned in a slow continuous motion several times. The transducer was held at various angles to display the maximum overall color signals throughout the three-dimensional circumference of the follicle wall.
For the spectral-Doppler mode, the setting for the range of detection of flow velocity was adjusted in each scanning to obtain the optimal spectral graph of velocities during a cardiac cycle [23]. The sample cursor or gate was set at a width of 1.5 mm, and the gate was placed on the image of the wall of the preovulatory follicle at the most prominent color signal that produced a spectrum of a cardiac cycle. A Doppler spectrum with three cardiac cycles (Fig. 1) was generated, and one of the cycles was used for velocity measurements. This was done three times, and the mean of the three measurements was used in the statistical analyses.
|
Oocyte Recovery and Processing
Oocyte recovery was done at Hour 30 by transvaginal ultrasound-guided aspiration of follicle contents and lavage with a 12-gauge double-lumen needle and vacuum pump (Fisher Scientific Inc., Pittsburgh, PA). The procedures for preparing the mare, transrectal ovary positioning, transvaginal transducer placement, inserting the needle, and the additives to the PBS (Sigma, St. Louis, MO) used for lavaging fluid have been described [1, 7]. Briefly, the follicular fluid was aspirated, and then the follicle wall of the aspirated follicle was lavaged with 180 ml of PBS per follicle. The lavaging was done 6–10 times, depending on follicle size. The aspirated follicular fluid and PBS lavaging fluid were held separately in a 37°C water bath until examined within 5 min after completing the recovery procedure. The follicular fluid was searched for an oocyte or cumulus-oocyte complex (COC). Then, the fluid was centrifuged (500 x g, 10 min), and 10 ml was stored at –20°C until hormone assay. The PBS lavaging fluid was filtered (70-µm BD Falcon Cell Strainer; BD Biosciences Discovery Labware, Bedford, MA), especially for passage and removal of red blood cells. The filter was rinsed, and the rinsings were placed in a Petri dish and searched with a stereomicroscope (Bausch & Lomb, Inc., New York, NY) for the oocyte or COC (if not found in the follicular fluid) and for granulosa cell evaluation. After evaluation of the COC, the cumulus was removed from the oocyte using 0.05% hyaluronidase and repeated pipetting. Sheets of mural granulosa cells were held by suction with a mouth pipette and washed three times in PBS. The sheets were fixed in 2% paraformaldehyde and stored at 4°C until evaluated for apoptosis. Apoptosis of the granulosa cells was used to indicate the maturity of the follicle [24, 25] and to evaluate the association of cell death with the oocyte recovery and maturity rates.
At both Hours 0 and 30, follicle diameter was obtained from the average of height and width of the antrum at the apparent maximal area from two frozen images. For color-Doppler mode at Hours 0 and 30, the percentage of circumference of the follicle wall with an apparent network of vessels was estimated in real time from the blood-flow color displays. Estimations of the proportion of the follicle circumference with blood-flow signals have been used in women [26] and mares [13, 14]. The percentage approach for the equine preovulatory follicle and corpus luteum has produced similar results by two operators working independently [14]. The velocity end points for the Doppler signals of a prominent vessel in the follicle wall were peak systolic velocity (PSV), time-averaged maximum velocity (TAMV), and end diastolic velocity (EDV). A drawing of a spectrum with the source for the velocity end points during a cardiac cycle is shown (Fig. 1). Resistance index (RI) and pulsatility index (PI) are computed from the velocities, as described [27]. A low RI or PI is related to low blood-flow impedance or high vascular perfusion in the tissues supplied by the artery distal to gate placement.
The imminence of ovulation at Hour 30 was evaluated in the preovulatory follicle in B mode as described [11]. The following discrete end points were recorded as present or absent: 1) serration of granulosa, 2) decreased turgidity, 3) loss of spherical shape, 4) apical area, and 5) echoic spots floating in the antrum. Serration of granulosa involved distinct irregularities or a notched appearance of both the inner and outer surfaces of the granulosum. Decreased turgidity was indicated when pressure was applied using the convex face of the transducer and was an indicator of an apparent decrease in intrafollicular pressure. Loss of spherical shape indicated that a spherical follicle at Hour 0 had become irregular in shape by Hour 30. An apical area was apparent when a formerly spherical follicle became smaller at one end (apex) and sometimes contained a stigma (nipple-like protrusion). Echoic spots in the follicular fluid floated in the antrum during ballottement.
The granulosa cells at Hour 30 were scored from 1–3 (minimal to maximal) for amount of cells and for the extent of expansion. Judging the amount of cells was aided by the extent of rinsing of the strainer as a result of clogging during passage of the lavaging fluid (score 1, minimal; score 2, 1 or 2 times; score 3, 3 times). Extent of cell expansion was estimated as follows: score 1, tight or compact; score 2, mixture of compact and expanded; score 3, expanded with a fluffy appearance. Apoptosis of the mural granulosa cells was evaluated by TUNEL assay (In situ cell death detection kit; Roche Diagnostic GmbH, Penzberg, Germany; [24]). The cells were processed according to the directions of the manufacturer of the TUNEL kit. After the TUNEL reaction, the cells were washed twice in PBS, and the nuclei of the cells were stained with DNA fluorochrome (10 µg/ml; Hoechst 33342; Sigma). A 10-µl cell suspension was placed on a slide and evaluated by fluorescent microscopy. At least 500 nuclei were counted for each sample. The percentage of apoptotic nuclei in the sample was used as the end point. The COC was classified as compact or expanded, as described [28]. The denuded oocyte was categorized as mature or immature on the basis of detection versus nondetection of a polar body, respectively. Categorization was limited to use of a stereomicroscope. Therefore, it was not determined whether oocytes that were categorized as immature were still in the germinal vesicle stage or in a progressive stage of meiosis that had not reached maturity (extruded polar body). The denuded oocytes were subjectively evaluated for cytoplasm morphology by using a stereomicroscope to estimate the percentage of ooplasm consisting of dark clusters of granules.
Estradiol was assayed in the plasma of heparinized jugular blood samples collected at Hours 0 and 30. Plasma concentrations of E2 were measured by a double-antibody radioimmunoassay kit, as described and validated in our laboratory for mare plasma [29]. The intraassay CV and sensitivity were 7.1% and 0.2 pg/ml, respectively. Estradiol, P4, testosterone, and free insulin-like growth factor (IGF) 1 were assayed in follicular-fluid samples collected at Hour 30. The concentrations of follicular-fluid hormones were determined by radio- or enzyme-immunoassays, as validated in our laboratory for equine follicular fluid [30]. The intraassay CV and the sensitivity for follicular-fluid E2 and testosterone, respectively, were 7.7% and 0.5 pg/ml, and 1.9% and 0.01 ng/ml. The intra- and interassay CV and sensitivity for follicular-fluid P4 and free IGF1 were 4.4%, 15.4%, and 0.06 ng/ml, and <1.0%, 9.4%, and 0.02 ng/ml, respectively.
Sequential data involving Hours 0 and 30 were analyzed by the SAS MIXED procedure to determine the main effects of group and hour and their interaction, using a Repeated statement to account for the autocorrelation between measurements (SAS version 8.2; SAS Institute Inc., Cary, NC). Student unpaired t-tests were used to locate differences between groups at each hour, and paired t-tests were used between hours within a group when a significant interaction or hour effect, respectively, was obtained. Differences between groups for end points available only at Hour 30 were analyzed by an unpaired t-test for quantitative end points and by chi-square for frequency end points. A probability of P < 0.05 indicated that a difference was significant, and P > 0.05 and 
0.1 indicated that significance was approached. Data are presented as the mean ± SEM unless otherwise specified.
An oocyte was recovered in 37 (62%) of 60 follicles. All oocytes were in the PBS lavaging fluid; none were in the aspirated follicular fluid. Follicle and hormone changes between the hour of hCG administration (Hour 0) and aspiration and lavaging of the follicle 30 h later (Hour 30) are shown (Fig. 2). The figure indicates means that were significant or approached significance for main effects of group (recovered versus nonrecovered oocyte) and hour and an interaction of group by hour. Both main effects and the interaction were significant for diameter; the interaction resulted from a smaller (P < 0.0009) diameter at Hour 30 for the group with recovered oocytes than for the group without recovered oocytes. For percentage of follicle circumference with blood-flow signals, both main effects approached significance from a tendency for increasing blood flow between hours (P < 0.07) and greater blood flow in the nonrecovery group (P < 0.08). A significant hour effect for TAMV, EDV, and RI resulted from an increase in blood velocity and a decrease in flow resistance, averaged for the two groups. The interaction for RI approached significance (P < 0.1), owing to a lower (P < 0.04) index at Hour 30 in the oocyte-recovery group. The results and statistical analyses for PI were similar to those for RI (PI data not shown). No significant effects were found for PSV (not shown). For plasma E2 concentrations, the main effect of hour (decrease in concentration) and the interaction were significant. The interaction resulted from greater concentration at Hour 0 that approached significance (P < 0.07) and lower (P < 0.03) concentration at Hour 30 in the recovery group than in the nonrecovery group.
|
The averages for characteristics available only at Hour 30 are shown, including the probabilities for a difference between groups (Table 1). For the discrete structural end points, the follicles with a recovered oocyte had a significantly greater frequency of serration and greater frequency of an apical area (approached significance). The score for amount of granulosa cells in the lavaging fluid and the score for granulosa expansion were greater for the follicles with a recovered oocyte. The frequency of apoptotic cells did not differ between groups. Concentrations of E2, P4, testosterone, and free IGF1 in the follicular fluid were not significantly different in the follicles with a recovered oocyte versus nonrecovered oocyte.
|
For the 37 recovered oocytes, 29 (78%) of 37 were mature (extruded polar body) and 22% were immature. Follicle and circulating E2 changes in the mature and immature groups between Hour 0 (hCG administration) and Hour 30 (recovery procedure) and the probabilities for differences are shown (Fig. 2). The main effect of hour and the interaction were significant for diameter; the interaction resulted from a smaller (P < 0.008) diameter at Hour 30 in the oocyte-mature group than in the immature group. For percentage of follicle circumference with blood-flow signals, the group effect (P < 0.06) and the hour effect (P < 0.08) approached significance, owing to a tendency for greater blood flow in the oocyte-mature group and an increase between hours averaged over the two groups; there was no interaction. No significant differences were found for PSV (not shown) and TAMV. For EDV, the group effect approached significance (greater in immature group; P < 0.09) and an increase between Hours 0 and 30, averaged over groups, resulted in a significant hour effect. An hour effect for RI resulted from a decrease but the group effect and interaction were not significant. Changes in PI and the statistical analyses (data not shown) were similar to those for RI. A decrease in plasma E2 between hours resulted in a significant main effect for hour. A significant interaction resulted from greater (P < 0.06) concentration at Hour 0 and lower (P < 0.009) concentration at Hour 30 in the oocyte-mature group than in the immature group.
Averages for follicle characteristics at Hour 30 for the groups with mature versus immature oocytes are shown, including the significant differences between groups (Table 1). The number of follicles with granulosa serration and the number with decreased turgidity were significantly greater in the group with mature oocytes. The scores for amount and expansion of granulosa, percentage of apoptotic cells, and frequency of follicles with an expanded COC were greater and percentage of ooplasm with dark clusters of granules was less in the group with mature oocytes. The follicles with a mature oocyte had significantly greater concentration of P4 and lower E2 and free IGF1.
The nuclear maturity of oocytes at Hour 30 was based on extrusion of the first polar body, whereas maturity of the ooplasm was based on the percentage of ooplasm with dark clusters. The changes in percentage of dark clusters in the ooplasm apparently involve the rearrangement of vesicles and lipid droplets from an even distribution to a more polarized appearance in mares [16]. The positive association between the percentage of ooplasm with dark granules and the oocyte maturation rate in the present in vivo study with equine oocytes is consistent with an in vitro study in cattle that indicated that oocytes with dark clusters of granules had a higher potential for maturation [31].
The higher percentage of granulosa cells with apoptosis in association with a mature oocyte supports, in vivo, a report [32] that the proportion of equine oocytes maturing in vitro was higher for the oocytes from follicles with apoptotic granulosa cells. In women, apoptosis has been found in follicles with oocytes that became fertilized [33]. It was proposed that apoptosis in the preovulatory follicle may be a normal physiologic process during luteinization of granulosa cells. Apoptosis of granulosa cells is also used as a marker of cell death during follicle atresia in various species, including equids [34]. In the present study, there was no indication of follicle atresia in that there was no detected effect of apoptosis on oocyte recovery rate.
Maturity of the follicle, as opposed to maturity of the oocyte, was judged by using discrete indicators of impending ovulation at Hour 30. Greater frequency of granulosa serration in both the recovery versus nonrecovery groups and in the oocyte-mature versus oocyte-immature groups indicated [11] that more mares in these groups had follicles that were close to ovulation. The low percentage of mares with development of an apical area or echoic spots in the antrum agrees with the previous findings [11] that these indicators of impending ovulation develop during the last few hours before ovulation. According to a previous study with a similar hCG protocol [13], most ovulations would be expected to occur before 18 hours after Hour 30. Greater maturity of the follicles with a recovered oocyte and nuclear mature oocyte also was indicated by the greater amount of granulosa in the lavaging fluid and greater expansion of the granulosa cells and COC [35, 36].
The change in follicle diameter during the 30 h following hCG treatment also was consistent with maturity of the follicle. In mares that are not treated with hCG, the follicle maintains an approximately constant diameter during the 36 h [13] or 2 days [37] before ovulation. In mares treated with hCG, the follicle did not increase in diameter before ovulation, whereas diameter increased in controls [13]. Thus, the smaller post-hCG diameter increase in the present study when an oocyte was recovered or the recovered oocyte was mature is attributable to greater maturity of the follicle by 30 h after hCG treatment.
The difference between groups in changes in systemic concentrations of E2 over Hours 0 to 30 are explainable on a similar basis as for diameter. Estradiol begins to decrease 2 days before ovulation [18]. The decrease in E2 between Hours 0 and 30 combined for all mares (main effect of hour) in the present results confirms the report that E2 decreases immediately after hCG is given [13]. The E2 results further support the conclusion of greater follicle maturity (closer to ovulation) at Hour 30 in the recovery and oocyte-mature groups. Presumably, therefore, preovulatory follicles at 32 mm differ considerably in the stage of maturity and in responsiveness to hCG.
Results on percentage of follicle circumference with blood-flow Doppler signals must be considered with reservation because the differences between groups and between hours only approached significance. A blood-flow increase between Hours 0 and 30 is consistent with a reported increase for 36 h after hCG treatment [13]. However, averaged over hours, blood flow was greater in the nonrecovery group than in the recovery group and greater in the oocyte-mature group than in the oocyte-immature group. This is inconsistent with the other results that indicated that the relationship between oocyte recovery rate and oocyte maturity rate was positive. For these reasons, the results on the effect of blood-flow signals on oocyte recovery and oocyte maturity are considered inconclusive. The spectral-Doppler blood velocities and indices did not show differences between the groups. Correction for the angle of impact of the ultrasound beams with blood flow (insonation angle) was not made because a defined and adequate vessel length was not available for placement of an angle cursor. Therefore absolute velocity values were not obtained, but the relative values among parts of the cardiac cycle are considered useful [23]. Combined for groups, the average relative velocity during a cardiac cycle (TAMV) and at the end of diastole (EDV) increased during the 30 hours, whereas the relative velocity at the peak of systole (PSV) did not change. The increase in relative TAMV and in EDV but not in PSV indicates that vascular adjustments were made during diastole rather than during systole. The continued flow of blood during diastole is a function of a rebounding effect of the arterial walls that were stretched during systole [23]. The present results indicate that elasticity of the arteries supplying the follicle increased between Hours 0 and 30.
The vascular perfusion indicators (RI and PI) showed a decrease in resistance to flow distal to the most prominent color signal in the follicular wall. In this regard, the RI taken at the most prominent signal in the preovulatory ovary was less for mares that became pregnant than for mares that did not [14]. In women treated for stimulation of multiple follicles, the RI and PI of an intraovarian artery were negatively correlated with the total number of retrieved oocytes [38], and recovery rates for mature and immature oocytes were greater for women with a low PI [39]. However, it does not seem appropriate to directly relate these previous findings in mares and women to those of the present study of a vessel in the wall of the solitary preovulatory follicle. In the present study, the increase in relative blood velocity during diastole and decrease in tissue resistance are both compatible with an expected requirement for an increased supply of blood as the follicle developed between Hours 0 and 30.
The absence of a difference in concentrations of follicular-fluid hormones between the oocyte-recovery and -nonrecovery groups, despite differences between the oocyte-mature and -immature groups, raises questions on the role of these hormones in follicle maturity versus oocyte maturity. The indicators of increased follicle maturity in the oocyte-recovery group (discussed above) were not accompanied by greater follicular-fluid concentrations of E2 and P4, and therefore did not show that these steroids were associated with follicle maturity. In contrast, mature oocytes compared to immature oocytes were associated with a greater follicular-fluid concentration of P4 and lower concentration of E2. In women, both E2 and P4 concentrations were greater in follicular fluid of follicles with mature versus immature oocytes [40, 41]. In a previous study in mares, E2 remained high throughout the oocyte maturation process, whereas P4 was higher during metaphase II than during earlier stages of nuclear maturation [16]. The present study provided temporal indications that intrafollicular E2 and P4 were not associated with follicle maturity but were associated with oocyte maturity or immaturity. This finding is novel but was unexpected and will require confirmation.
The associations between intrafollicular versus plasma concentrations of P4 and testosterone are of interest. Plasma P4 does not begin to increase until 12 h after ovulation [19], yet follicular-fluid P4 increased dramatically before ovulation (20-fold; [15]) and in the oocyte-mature group of the present study. Testosterone concentrations in follicular fluid were not different between groups at Hour 30 but reportedly increase during the preovulatory period in both the follicular fluid [15] and in the plasma [18]. These considerations are compatible with the present interpretation of an intrafollicular role of P4 in oocyte maturation and a reported systemic effect of testosterone on accumulation of FSH in the pituitary during estrus in mares [42]. Thus, the effects of follicle-produced P4 may be local, as opposed to only systemic effects of testosterone.
There was an indication that the bioavailability of IGF1 was reduced during the oocyte maturation process in the sense that the concentration of free IGF1 in the follicular fluid was lower in the oocyte-mature group than in the oocyte-immature group. The higher concentrations of free IGF1 and lower frequency of apoptotic granulosa cells in follicles with immature oocytes may be related in that IGF1 is an inhibitor of apoptosis in many types of cells [43]. The present IGF1 finding in mares is consistent with the reports in women that concentrations of IGF binding proteins were higher in the follicular fluid from follicles with mature oocytes than with immature oocytes [40, 41]. Furthermore, IGF binding protein levels were positively correlated with levels of E2 and P4 but not with androstenedione [41], which is compatible with the present findings in equine follicles with mature oocytes. The role of an in vivo reduction in bioavailability of IGF1 during oocyte maturation apparently is not known. Although the results of the present in vivo study with equine follicles indicated that IGF1 concentrations were lower in follicles with a mature oocyte than with an immature oocyte, in vitro studies in many species have indicated that IGF1 enhances the nuclear and cytoplasmic maturation of oocytes [44, 45]. During in vitro maturation of equine oocytes, maturation increased when the medium contained IGF1 alone but not when FSH, LH, E2, and fetal calf serum were added [46]. Further study and clarification is needed on the role of IGF1 and its binding proteins on in vivo maturation of the oocyte.
The results of the present study have comparative relevance, owing to the similarities in dynamics of the ovulatory follicular wave between mares and women [47]. The similarities include an approximately twofold greater diameter of the largest follicle in mares than in women, extending from diameter at the peak of the follicular wave-inducing FSH surge until preovulatory diameter is maximum. In addition, hCG is used to induce final follicle and oocyte maturation and ovulation in both mares [48] and women [49, 50]. The diameter of the dominant follicle at the time of treatment (mares, 32–36 mm; women, 16–18 mm) conforms with the twofold difference between species in follicle diameter, and the interval from treatment to ovulation (36–48 h) is similar.
In conclusion, the results indicated that recovery of an oocyte and maturity of the recovered oocyte are both dependent upon the rate of follicle maturity in response to the hCG treatment. Our clinical interpretation is that the success of recovery attempts that are made on the basis of number of hours after hCG treatment is influenced by the wide variation in the response to treatment. Results also showed that follicular-fluid concentrations of P4 were higher, E2 and free IGF1 were lower, and testosterone did not change in the oocyte-mature group compared to the oocyte-immature group, but that concentrations of these hormones were not different between the oocyte-recovery and nonrecovery groups. Our physiologic interpretation on a temporal basis is that follicular-fluid P4 and the IGF system are associated with final oocyte maturation but not with follicle maturation.
ACKNOWLEDGMENTS
The authors thank J.-C. Ju for advice and technical assistance and Dee Cooper for assistance with statistics and manuscript preparation.
FOOTNOTES
1Supported by the Eutheria Foundation (Cross Plains, WI). Project P2-OG-06. 
Correspondence: 2O.J. Ginther, Department of Pathobiological Sciences, School of Veterinary Medicine, 1656 Linden Drive, University of Wisconsin, Madison, WI 53706. FAX: 608 262 7420; e-mail: ginther{at}svm.vetmed.wisc.edu
Received: 28 February 2007.
First decision: 28 March 2007.
Accepted: 26 April 2007.
REFERENCES
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
