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a Department of Biology, Trinity University, San Antonio, Texas 78212
b Department of Cell Biology, Baylor College of Medicine, Houston, Texas 77030
c Department of Gynecology and Obstetrics, Kyoto University Medical School, Kyoto, Japan
d Department of Obstetrics and Gynecology, Kumamoto University School of Medicine, Kumamoto, Japan
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-OH progesterone decrease in the ovary before the follicle ruptures, while progesterone, prostaglandin E2, and prostaglandin F2 increase enormously during the ovulatory process [1]. The regulation of these follicular steroids and prostanoids by various enzymes and enzyme inhibitors has been studied extensively during the past three decades [2,3]. However, there is only limited information on the relationship of these ovarian compounds to gonadotropin-induced ovarian carbonyl reductase activity during ovulation. Evidence exists that several isoenzymes of carbonyl reductase increase in the ovary in response to various dosages of gonadotropin(s) [4–9], and there is more recent evidence of ovarian expression of several homologous genes for carbonyl reductases [10,11]. Yet the ovarian functions of these aldo-keto reductases are not clearly understood. In our recent use of differential-display polymerase chain reaction [12] to identify ovarian gene expression during ovulation, we isolated four slightly different isoforms of genes for carbonyl reductases on four different occasions (i.e., using four different primer sets for the reverse transcription-polymerase chain reaction [RT-PCR] reaction in the differential-display procedure). Northern blotting has confirmed that these carbonyl reductase genes are distinctly expressed in the rat ovary during ovulation. The purpose of this study was to investigate the temporal expression, cellular distribution, and regulation of carbonyl reductase gene expression during the ovulatory process in the gonadotropin-primed immature rat model.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Immature Wistar rats were selected from litters in a breeding colony consisting of 80 adult females and 40 males. Animals were housed at 23°C under 14L:10D (lights-on at 0700 h). Immature rats were chosen because in this experimental model it is easy to induce superovulation and extirpate ovaries that consist mainly of gonadotropin-responsive follicular tissue. In addition, ovaries of immature rats do not contain the atretic follicles or corpora lutea characteristic of cycling adult animals. Therefore, the ovarian response to gonadotropic stimulation can be attributed primarily to preovulatory follicles. For the determination of ovulation rate and the extraction of ovarian RNA, rats were killed by exposure to CO2. The animals were acquired, retained, and used in compliance with the NIH Guide and with the approval of the Institutional Animal Care Review Board.
Ovulation Induction
The young females in the litters were weaned at ~23 days of age. The animals selected for experimentation were within the weight range of 40–50 g. From our experience, body weight is more important than age for obtaining a consistent ovulation rate upon stimulating animals with specific doses of gonadotropins (unpublished results). At 0800–0900 h on the day after weaning, the immature rats were injected s.c. with 10 IU eCG (G4877; Sigma Chemical Co., St. Louis, MO). At 0800–0900 h, 2 days later, the ovulatory process was initiated by 10 IU hCG (CG-5; Sigma) s.c. Using this protocol, the animals begin ovulating approximately 12 h after the hCG injection. At 24 h after hCG was injected, ovulation rate was determined by compressing the oviducts between two microscope slides and counting the number of ova that had accumulated in the oviducts. The specified doses of eCG and hCG yield approximately 60–70 mature follicles per pair of ovaries (i.e., per animal), and approximately 60–70% of the ovarian mass consists of mature follicles. In the experiments to test the effects of epostane and indomethacin on ovulation rate, eight rats were used in each control and each treated group of animals.
Epostane and Indomethacin Administration
Epostane (courtesy of Sterling-Winthrop Research Institute, Rensselaer, NY) was suspended in distilled water, and 1.0 N NaOH was added drop-wise during vigorous stirring until the agent dissolved. The pH of the epostane solution was adjusted to 7.8–8.0 by adding 0.1 N HCl during stirring. This synthetic steroid derivative was given s.c. in a dose of 5.0 mg/rat at 3 h after hCG was administered. Indomethacin (I7378; Sigma) was dissolved in sodium carbonate solution containing 4 mg of Na2CO3 for every 10 mg of indomethacin. After ~15 min of stirring, the indomethacin solution was adjusted to pH 7.4–7.6 by slowly adding 0.1 N HCl during vigorous stirring. This nonsteroidal anti-inflammatory drug was given s.c. in a dose of 1.0 mg/rat at 3 h after administration of hCG to induce the ovulatory process. Ovarian RNA was extracted for Northern blotting according to the procedure described below. Control RNA was extracted from five ovaries each at 0 h and 8 h after the administration of hCG, and RNA from animals treated with epostane or indomethacin were extracted from five pairs of ovaries each at 8 h after induction of the ovulatory process with hCG.
RNA Extraction
Total RNA was extracted from whole ovaries that were extirpated at 0, 2, 4, 8, 12, and 24 h after the ovulatory process had been initiated by a dose of 10 IU of hCG. (Since follicles begin rupturing approximately 12 h after exposure to exogenous hCG, the six different extracts of RNA represented one preovulatory control group, four experimental groups taken at the specified intervals during the ovulatory process, and one postovulatory control group that contained newly developing corpora lutea.) At each of the six designated intervals after hCG, the ovaries from 12–20 rats were immediately frozen in liquid nitrogen and then pooled together to provide a total of approximately 1 g of ovarian tissue for RNA extraction based on the procedure of Chomczynski and Sacchi [13]. Each gram of ovarian tissue was placed in 3.0 ml of 4 M guanidine isothiocyanate solution and homogenized for 20–30 sec using a Tissue Tearor homogenizer (985-370; Biospec Products, Bartlesville, OK). The homogenized samples were layered onto 1.5 ml of 5.7 M cesium chloride and centrifuged overnight at 126 000 x g. After extraction in phenol/chloroform/isoamyl alcohol, the total RNA in each extract was diluted 100-fold and quantitated at 260 and 280 nm to estimate the concentration of RNA in the original extract and to be sure there was not excessive protein in the samples. To reduce the amount of DNA in the nucleic acid extracts prior to the RT-PCR reactions, 50-µg aliquots of RNA were exposed to DNase from a MessageClean kit (M601; GenHunter Corporation, Nashville, TN) available for this specific purpose. After this cleaning step, the RNA was quantitated by spectrophotometry once again, and aliquots were diluted to a concentration of 0.1 µg RNA/µl H2O for use in the RT step as described below.
RT and PCR Amplification
The RT-PCR reactions for differential display were carried out using RNAimage kits (G501, G508; GenHunter). Briefly, for RT, 0.2 µg of the clean total RNA was primed with one-base-anchored oligo(dT) primers containing a HindIII site (H = AAGCTT) attached to the 5'-end (i.e., 5'-HTTTTTTTTTG-3' and 5'-HTTTTTTTTTA-3'), and the reaction was carried out with Moloney murine leukemia virus reverse transcriptase. The thermocycler (Thermolyne Temp Tronic model #DB66925; Barnstead/Thermolyne Corp., Dubuque, IA) was programmed at 65°C for 5 min, 37°C for 60 min, and 95°C for 5 min, followed by a 4°C hold. The first-strand cDNAs were amplified subsequently by PCR using selected 13-mer arbitrary upstream primers (as designated at the beginning of Results) from the two RNAimage kits in conjunction with the oligo(dT) primers. The amplification was carried out with Taq DNA polymerase (M1861; Promega, Madison, WI), and the reaction mixture included [35S]dATP (NEG034H; NEN Life Science Products, Boston, MA) for labeling the PCR products. The thermocycler was programmed for an initial 2-min denaturation at 94°C, followed by 40 cycles of amplification with denaturation at 94°C for 30 sec, annealing at 38°C for 2 min, and extension at 72°C for 30 sec, followed by a 72°C dwell for 5 min and a 4°C hold.
Electrophoresis to Detect Differentially Expressed cDNA
The PCR products were separated by electrophoresis on a sequencing gel consisting of 6% acrylamide and 48% (w:v) urea in single-strength TBE buffer (89 mM Tris-borate, 2 mM EDTA). Before loading onto the gel, 3.5 µl of each PCR sample was mixed with 2.0 µl of loading dye and heated at 80°C for 3 min. Electrophoresis was carried out at a gel temperature of 50–55°C. The gels were dried onto Whatman (Clifton, NJ) 3M paper and then stapled to HyperFilm ßMAX (RPN11; Amersham Pharmacia Biotech, Piscataway, NJ) for 1–3 days. Bands of cDNA that were differentially displayed during the six time intervals of the periovulatory period were visually selected for further analysis.
Extraction and Reamplification of Potentially Unique cDNA
The unique cDNA bands that were identified on the autoradiographs were located in the acrylamide gel (on Whatman paper) by aligning the staple holes in the paper with the staple holes in the corresponding autoradiograph. After comparison of six parallel lanes of electrophoretically separated cDNA, only the single most intense band was excised from the dried gel. The excised band was rehydrated in 100 µl of H2O and boiled for 15 min before the cDNA was recovered by ethanol precipitation. The recovered cDNA was reamplified using the same original primer set and initial PCR conditions, except that the dNTP concentration in the reaction mixture was 20 µM and radioactive dATP was not added. Before this PCR product was used as a probe for Northern blot analysis, the size, purity, and amount of the amplified cDNA was estimated by electrophoresis of a 10-µl aliquot of the sample on a 2% agarose (US32829; Amersham) minigel. The cDNA was visualized by ethidium bromide staining, and the size and amount of each PCR product were estimated through correlation to a ladder of standards consisting of 50, 150, 300, 500, 750, and 1000 base pairs (bp) (G3161; Promega). If the amount of cDNA in the 10-µl aliquot of PCR product was equal to or greater than the amount in the bands of the standards (i.e., 
300 ng), then the PCR product was used for Northern blotting. However, if the PCR product band was less intense than the nearest band of cDNA standard, then the first-round PCR sample was diluted 1:10, and 4 µl of this dilution was used as the template for another 40-cycle PCR amplification before Northern blotting was performed.
Preliminary Probing of Northern Blots with Selected cDNA Fragments
Minigels consisting of 1.2% agarose (US75817; Amersham) in 6.7% formaldehyde and 1 M Hepes/NaPO4 (pH 7.0) were used for electrophoresis of the total RNA to prepare Northern blots. Aliquots containing 20 µg of the original total RNA extracted from each of the six stages of ovaries were loaded into adjacent wells of a minigel, and electrophoresis was carried out at 38 V. After electrophoresis, the RNA in the minigel was blotted onto a nylon membrane by standard procedures, and the Northern blot was vacuum baked at 80°C for 100–120 min. Radiolabeling of the PCR product (i.e., of the differential-display amplicon) with [32P]dCTP (NEG013H; NEN) was facilitated by a Prime-a-Gene Labeling System (U1100; Promega). Unincorporated dNTPs were removed from the labeling mixture by centrifugation through NICK-Spin Columns (17-0862; Amersham). The hybridization mixture contained approximately 2 000 000 cpm of probe/1.0 ml of solution. After 15–18 h of hybridization, the blots were rinsed to a stringency of 0.5-strength SSC (single-strength SSC is 0.15 M sodium chloride and 0.015 M sodium citrate)/0.1% SDS at 55°C and exposed to Hyperfilm MP (RPN6H; Amersham) for an appropriate amount of time. To confirm that the different lanes on the Northern blots had equivalent amounts of RNA, the blots were stripped and then reprobed with ß-actin cDNA (7323; Ambion, Austin, TX).
Densitometric Analysis of Northern Blots
The density of the signals from the Northern blots was analyzed by the NIH-image program (http://rsb.info.nih.gov/nih-image/) using the download "NIH-image 162 fat.hqx." Each Northern analysis of a uniquely expressed cDNA was scanned simultaneously with the ß-actin control for the same Northern, and the image was saved in Adobe Photoshop (Mountain View, CA). The corresponding lanes on the experimental and control blots were aligned, and the image was rotated 90° in order to use the rectangular selection tool of the NIH-image program to select an area that encompassed each cDNA experimental lane with its corresponding ß-actin control lane. After digitizing all the bands on the Northerns, the ratio of the density of each experimental band to its corresponding ß-actin control band was calculated for each lane on the Northerns. These ratios having been obtained, the value of the 8-h lane was arbitrarily established as 100%, and the densities of all of the other lanes were expressed as percentages of the 8-h value.
Cloning of Potentially Unique cDNA
On the basis of the Northern blot patterns, cDNA fragments that appeared to be unique to ovulation were cloned. This step was facilitated by a pCR-TRAP Cloning System (P404; GenHunter). PCR products from reamplification of a potentially unique cDNA were blunt-end ligated into a specific cloning site of the plasmid vector that endowed competent Escherichia coli with tetracycline resistance. The E. coli were grown on LB agar plates that contained 10 µg/ml tetracycline. To identify a colony that contained plasmids with the desired cDNA insert (rather than plasmids with stray nucleotides or primers), 6–8 colonies were scored and small amounts of the colony cells were transferred by a pipette tip to a microfuge tube containing lysis buffer. The cDNA inserts were PCR amplified out of each of the colony plasmids using a primer set (available in the pCR-TRAP Cloning System) that flanked each side of the plasmid insert site by 62 bp. Subsequently, the sizes of the PCR products were analyzed by electrophoresis on a 2% agarose minigel containing ethidium bromide for visualization of the cDNA. Only amplified cDNAs of a size comparable to the size initially estimated at the time of extraction from the differential-display gel were analyzed further. (In this instance, the cDNA fragments on the minigel were selected only if they were the original size plus 124 bp, which takes into account the flanking segments of plasmid DNA that were amplified simultaneously with the insert.)
Confirmation of E. coli Colonies with Inserts of Unique cDNA
When a potentially unique amplicon was originally cut out of the differential-display gel, it was likely that the extirpated Whatman 3M paper contained additional cDNA fragments of approximately the same size as the unique cDNA. Therefore, although a given colony might have contained a cDNA insert that appeared to be the appropriate size, the insert was not necessarily the same cDNA fragment that exhibited differential hybridization during the initial Northern blot analysis. To identify colonies that actually contained cDNA fragments unique to ovulation, aliquots of all of the PCR samples from the previous step that contained the predicted size of cDNA insert were used to probe additional Northern blots. This secondary analysis by Northern blotting reconfirmed the uniqueness of a given cDNA fragment while simultaneously identifying specific cloning colonies containing the unique cDNA.
Sequencing the Unique cDNA Fragments
Samples of colonies containing unique cDNAs were cloned further in LB medium containing 10 µg tetracycline/1.0 ml medium. The harvested plasmid DNA was purified using a Wizard Minipreps DNA Purification System (A7500; Promega). The double-stranded plasmid DNA was prepared for sequencing by an AidSeq Kit C (P203; GenHunter). Manual sequencing was performed using a Sequenase Version 2.0 DNA Sequencing kit (US70770; Amersham) utilizing [35S]dATP (NEG034H; NEN). The nucleic acid sequences were analyzed by a BLASTn search of the database server at the National Center for Biotechnology Information (NCBI) (http://www.ncbi.nlm.nih.gov).
In Situ Hybridization
Ovaries were removed from three individual rats at each of the six periovulatory intervals described in the section on RNA extraction and were fixed overnight in 4% paraformaldehyde. The in situ hybridization protocol was adapted from a published method [14]. The procedure was facilitated by using a Riboprobe Transcription System (P1450; Promega).
Statistical Analysis
Numerical data are presented as means ± SE. The significances of the differences were determined by Duncan's Multiple Range tests after a completely randomized one-way ANOVA of the means of the groups. The probability value used as the cut-off between "significant" and "not significant" was P = 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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During this study, 144 different primer sets were used in the PCR reactions to amplify the cDNA generated from the ovarian mRNA extracted at the 6 designated intervals during the periovulatory period. Four of those primer sets resulted in differential-display autoradiographs that revealed ovulation-specific expression of cDNAs associated with carbonyl reductase (Fig. 1). The particular pairs of primers that yielded differentially expressed cDNAs homologous to carbonyl reductase genes were 1) 5'-HTTTTTTTTTG-3' and 5'-HCTCAACG-3' (H-AP4; GenHunter); 2) 5'-HTTTTTTTTTG-3' and 5'-HAACGAGG-3' (H-AP7); 3) 5'-HTTTTTTTTTG-3' and 5'-HAACTGAG-3' (H-AP58); and 4) 5'-HTTTTTTTTTA-3' and 5'-HAACTGAG-3' (H-AP58). The differentially expressed cDNAs are designated as G4, G7, G58, and A58, respectively, on the basis of the primer sets that amplified them (Fig. 1).
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Northern Blots of Carbonyl Reductase mRNA Expression During Ovulation
Each of the cDNA fragments generated by the four sets of primers hybridized in a similar pattern to Northern blots. The cDNA probes generated minimal signals in the 0-h and 24-h lanes of the Northerns (Fig. 2), while the strongest signal was at 8 h after hCG. The strongest signal (i.e., the 8-h lane) was arbitrarily set at 100%, and the densities at the other intervals of the ovulatory process were expressed as fractions of that maximum. Accordingly, the signal densities (normalized against the ß-actin control) at 0, 2, 4, 8, 12, and 24 h after hCG were 7.4%, 25.9%, 71.9%, 100%, 57.2%, and 8.2%, respectively (Fig. 2). Thus, during the first 8 h of the ovulatory process, there was a 14-fold increase in carbonyl reductase gene expression. Subsequently it declined, resulting in expression at 24 h that was not significantly different from the 0-h control level.
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Sequences of the cDNA Fragments for Carbonyl Reductase
Sequencing revealed that the cDNA fragments differentially displayed in Figure 1 were all approximately 300 bp in length (Fig. 3). However, there were slight variations in the lengths of the cDNAs, for three distinctly different reasons. First, as expected, the number of base pairs in a given fragment varied depending on the site where the random primer annealed to the first strand cDNA (Fig. 3). Second, the poly-T primers exhibited slight variations in the manner in which they annealed to the mRNA (or the cDNA). Third, four of the eight different clones that were sequenced had a 7-bp deletion located approximately one third of the distance from the poly-T tail. In addition, there were four other positions along this ~300-bp segment of carbonyl reductase genes that exhibited polymorphism (Fig. 3). Close comparison of the sequences of the eight different clones revealed that they were fragments of four different forms of an ovarian carbonyl reductase gene. Clones G7-1 and G4-1 (NCBI Accession #AF181955) were essentially the same except for a slight variation in the way the poly-T primer annealed to the fragment (Fig. 3). Clones A58-1 and G58-1 (NCBI Accession #181956) were identical also, except for a slight difference at the poly-T end. Clones G4-2 and G7-2 (NCBI Accession #AF023086) were basically identical to one another, except for the 3-bp difference in the site of attachment of the random primer. Clones G7-3 and G7-4 (NCBI Accession #AF181957) were identical to one another. Furthermore, the existence of these four polymorphs explains why, in at least one instance, two different lengths of unique cDNAs were visible in the same differential-display autoradiograph. Specifically, in Figure 1, the autoradiograph corresponding to the G7 primer set reveals a longer cDNA (i.e., G7-2) and a shorter cDNA (i.e., G7-1) that are both uniquely expressed during ovulation. Also, sequence analyses of several different clones from the slightly distinguishable double band in the G4 autoradiograph (Fig. 1) verified that at least two of the three different cDNA polymorphs were produced by this primer set. The sequences of all eight clones were confirmed by sequencing the full length of each cDNA with both a forward and a reverse primer, i.e., by sequencing both strands of each cDNA.
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Localization of Carbonyl Reductase mRNA Expression by In Situ Hybridization
In view of the degree of homology among the eight cDNAs, only one (i.e., the 305-bp G7-2 fragment) was used for in situ hybridization analysis. The results confirmed the temporal pattern of carbonyl reductase mRNA expression observed by Northern blotting. There was very limited signal from the probe at 0 h and 24 h into the ovulatory process, while the strongest signal was at 8 h after the rats were stimulated by hCG (Fig. 4). Hybridization was intense in the thecal layers of mature follicles, as well as in extrafollicular connective tissue throughout the ovary, while there was no significant hybridization in the granulosa layer (Fig. 5).
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Effects of Epostane and Indomethacin on Carbonyl Reductase Gene Expression
For these tests, Northern blots were prepared from RNA that had been extracted from control ovaries at 0 and 8 h into the ovulatory process and from experimental ovaries taken at 8 h after hCG from rats treated 5 h earlier with ovulation-inhibiting doses of epostane or indomethacin. As in the Northern blotting tests (described above) at the six different intervals during ovulation, the signal density (normalized against the ß-actin control) of the 8-h lane was arbitrarily set at 100% (Fig. 6). This intensity at 8 h was 13-fold greater than the 0-h control value for carbonyl reductase gene expression. In animals treated with the anti-ovulatory agent epostane, which blocks progesterone synthesis, the signal density level of 110.6% was not significantly different from the 8-h control value. Similarly, indomethacin, which blocks prostanoid synthesis, resulted in a signal density level of 120.5%, which was not significantly different from the 8-h control value. Thus, neither of these two well-known ovulation-inhibiting agents reduced the ovarian expression of carbonyl reductase genes.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The G4-1/G7-1 cDNA fragments are identical to segments of a 981-bp mRNA (NCBI Accession #D89069) that is reportedly inducible in immature rat ovaries that have been stimulated by eCG alone [11]. Similarly, the G58-1/A58-1 cDNA fragments are identical to a 997-bp mRNA (NCBI Accession #X84349) found in rat testes [17]. In contrast, the G4-2/G7-2 and the G7-3/G7-4 cDNA isoforms of a carbonyl reductase gene are not identical to any other subjects registered in the NCBI gene database. On the other hand, all eight of the clones are highly homologous to a 1053-bp mRNA (NCBI Accession #U31966) for carbonyl reductase in the mouse cerebellum [18]. Likewise, they are similar to a segment of a noninducible carbonyl reductase gene (NCBI Accession #D89070) in the rat ovary [11]. Thus, the cDNA fragments presently discovered by the randomness of the differential-display procedure are polymorphic representations of the superfamily of genes that translate into aldo-keto reductases associated with steroid and eicosanoid metabolism in a number of tissues.
The physiological role of these aldehyde- and ketone-reducing enzymes is not clear [17]. They were first associated with ovarian function a decade ago [4]. However, subsequent efforts to clarify their role in ovarian follicular events have led to confusing conclusions. Diverse results have suggested that ovarian gene expression for carbonyl reductase is important "in ovarian follicular development" [11], but "is not always necessary for follicular maturation" [9]. In similar contradiction, other reports conclude that carbonyl reductase "is closely involved . . . in the ovulatory process" [6], but "is not always necessary for . . . the ovulatory process" [8]. It has been further suggested that, rather than contributing to follicular maturation or ovulation, carbonyl reductase may be "involved in the formation of neo corpora lutea after ovulation" [8]. This uncertainty about the function of ovarian carbonyl reductase may be the consequence of use of a variety of unusual protocols for treating rats with gonadotropins in diverse studies during the past decade [4–9,11]. On the other hand, the present study provides the first clear analysis of the temporal pattern of expression of four polymorphs of ovarian carbonyl reductase genes during ovulation in the common experimental model of the gonadotropin-primed immature rat.
The present study also demonstrates that the temporal pattern of expression of carbonyl reductase mRNA in eCG-primed immature rats is the same, regardless of whether one examines the differential-display gels, the Northern blots, or the in situ hybridization data. There was a slight increase in carbonyl reductase mRNA at 2 h after induction of ovulation by treatment with hCG. The expression reached a peak at 8 h after hCG, and it remained relatively high at 12 h after hCG, i.e., when follicles first begin to rupture. Subsequently, the ovarian mRNA for carbonyl reductase returned to control levels by 24 h after hCG, i.e., by 10–12 h after completion of the ovulatory process (an observation that does not support a role for this enzyme in luteal development). This postovulatory down-regulation is not consistent with earlier reports [4,8] that ovarian carbonyl reductase enzyme activity is the highest after ovulation, i.e., in neonatal tissue. This discrepancy may be due to some difference between the temporal pattern of mRNA expression versus the half-life of the proteinaceous enzyme.
This report contains the first in situ hybridization data on the localization of ovarian carbonyl reductase mRNA expression. Most of the signal was from the thecal layers of the larger follicles, along with additional expression in the extrafollicular connective tissue in the ovary. In contrast, there was negligible expression of carbonyl reductase gene in the granulosa layer of the follicles. These observations are in spatial (but not temporal) conformity with earlier immunohistochemical data that localized the actual enzyme in ovarian thecal tissue [4].
It remains to be determined why carbonyl reductases are expressed so strongly in ovarian thecal connective tissue during ovulation. The temporal pattern of their expression suggests that they must be related in some way to the ovulatory process. However, it is not possible to establish from the present data that carbonyl reductase activity contributes in any direct way to the mechanism of ovulation, because doses of epostane and indomethacin that inhibited ovulation did not block the transcription of mRNA for this enzyme. This finding suggests that neither progesterone nor prostanoids are required for the induction of ovarian carbonyl reductase expression. Yet it is not possible to exclude a role for this enzyme in the ovulatory process, because the gonadotropin-induced expression of carbonyl reductase could precede the increases in ovarian steroids and prostanoids within the chronology of ovarian events that lead to follicular rupture. Conversely, since neither epostane nor indomethacin completely blocks the ovulatory increase in progesterone [21,22], it is still possible that this steroid may be associated with carbonyl reductase gene expression in the ovary.
Finally, studies in other biological systems have led to the hypothesis that aldo-keto reductases may have a physiologic role in the detoxification of steroids, prostanoids, and pterins in tissues where their endogenous production is excessive [16,20,23]. Therefore, it is possible that carbonyl reductase may be serving as a local protective response to the substantial increases in ovarian steroids and prostanoids during ovulation. It would be interesting to know whether the periovulatory expression of this enzyme might also influence the mobilization of follicular fibroblasts that is known to occur at the time of ovulation. In any event, the significant expression of carbonyl reductase mRNA in the fibroblast-rich thecal layers of the follicle suggests that ovarian carbonyl reductases have some significant ovarian function at the time of ovulation. Their potential influence on thecal fibroblasts during ovulation is worthy of further analysis.
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FOOTNOTES |
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1 Supported by NSF grant #9870793 (L.L.E.), by a grant to support T.U. as a Research Fellow of The Lalor Foundation, Providence, RI (L.L.E.), and by NIH grant HD-16229 (J.S.R.). This work was presented in part at the 30th Annual Meeting of The Society for the Study of Reproduction held at Portland, OR, August 2-5, 1997. 
2 Correspondence. FAX: 210 999 7229; lespey{at}trinity.edu
Accepted: September 9, 1999.
Received: June 17, 1999.
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during ovulation in the gonadotropin-primed immature rat. Endocrinology 1990; 127:259–263.This article has been cited by other articles:
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B. Senthilkumaran, C.-C. Sudhakumari, X.-T. Chang, T. Kobayashi, Y. Oba, G. Guan, Y. Yoshiura, M. Yoshikuni, and Y. Nagahama Ovarian Carbonyl Reductase-Like 20{beta}-Hydroxysteroid Dehydrogenase Shows Distinct Surge in Messenger RNA Expression During Natural and Gonadotropin-Induced Meiotic Maturation in Nile Tilapia Biol Reprod, October 1, 2002; 67(4): 1080 - 1086. [Abstract] [Full Text] [PDF]
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M. Tanaka, S. Nakajin, D. Kobayashi, S. Fukada, G. Guan, T. Todo, B. Senthilkumaran, and Y. Nagahama Teleost Ovarian Carbonyl Reductase-Like 20{beta}-Hydroxysteroid Dehydrogenase: Potential Role in the Production of Maturation-Inducing Hormone During Final Oocyte Maturation Biol Reprod, May 1, 2002; 66(5): 1498 - 1504. [Abstract] [Full Text]
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L. L. Espey, S. Yoshioka, T. Ujioka, S. Fujii, and J. S. Richards 3{{alpha}}-Hydroxysteroid Dehydrogenase Messenger RNA Transcription in the Immature Rat Ovary in Response to an Ovulatory Dose of Gonadotropin Biol Reprod, July 1, 2001; 65(1): 72 - 78. [Abstract] [Full Text] [PDF]
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S. Yoshioka, S. Ochsner, D. L. Russell, T. Ujioka, S. Fujii, J. S. Richards, and L. L. Espey Expression of Tumor Necrosis Factor-Stimulated Gene-6 in the Rat Ovary in Response to an Ovulatory Dose of Gonadotropin Endocrinology, November 1, 2000; 141(11): 4114 - 4119. [Abstract] [Full Text] [PDF]
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T. Ujioka, D. L. Russell, H. Okamura, J. S. Richards, and L. L. Espey Expression of Regulator of G-Protein Signaling Protein-2 Gene in the Rat Ovary at the Time of Ovulation Biol Reprod, November 1, 2000; 63(5): 1513 - 1517. [Abstract] [Full Text]
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L. L. Espey, T. Ujioka, D. L. Russell, M. Skelsey, B. Vladu, R. L. Robker, H. Okamura, and J. S. Richards Induction of Early Growth Response Protein-1 Gene Expression in the Rat Ovary in Response to an Ovulatory Dose of Human Chorionic Gonadotropin Endocrinology, July 1, 2000; 141(7): 2385 - 2391. [Abstract] [Full Text] [PDF]
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E. Brockstedt, M. Peters-Kottig, V. Badock, C. Hegele-Hartung, and M. Lessl Luteinizing Hormone Induces Mouse Vas Deferens Protein Expression in the Murine Ovary Endocrinology, July 1, 2000; 141(7): 2574 - 2581. [Abstract] [Full Text] [PDF]
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L. L. Espey, S. Yoshioka, D. L. Russell, R. L. Robker, S. Fujii, and J. S. Richards Ovarian Expression of a Disintegrin and Metalloproteinase with Thrombospondin Motifs During Ovulation in the Gonadotropin-Primed Immature Rat Biol Reprod, April 1, 2000; 62(4): 1090 - 1095. [Abstract] [Full Text]
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