Biol Reprod Keystone Symposia Conference on Frontiers in Reproductive Biology & Regulation of Fertility.
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Biology of Reproduction 66, 1413-1421 (2002)
© 2002 Society for the Study of Reproduction, Inc.

Regular Article

Gonadotropin Surge-Induced Up-Regulation of the Plasminogen Activators (Tissue Plasminogen Activator and Urokinase Plasminogen Activator) and the Urokinase Plasminogen Activator Receptor Within Bovine Periovulatory Follicular and Luteal Tissue1

Mark P.D. Dowa,b, Leanne J. Bakkeb, Carolyn A. Cassara, Michael W. Petersa, J. Richard Pursleya, and George W. Smith2,,a,b

a Departments of Animal Science b Physiology, Michigan State University, East Lansing, Michigan 48824-1225


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
This study examined the effect of the preovulatory gonadotropin surge on the temporal and spatial regulation of tissue plasminogen activator (tPA), urokinase plasminogen activator (uPA), and uPA receptor (uPAR) mRNA expression and tPA, uPA, and plasmin activity in bovine preovulatory follicles and new corpora lutea collected at approximately 0, 6, 12, 18, 24, and 48 h after a GnRH-induced gonadotropin surge. Messenger RNAs for tPA, uPA, and uPAR were increased in a temporally specific fashion within 24 h of the gonadotropin surge. Localization of tPA mRNA was primarily to the granulosal layer, whereas both uPA and uPAR mRNAs were detected in both the granulosal and thecal layers and adjacent ovarian stroma. Activity for tPA was increased in follicular fluid and the preovulatory follicle apex and base within 12 h after the gonadotropin surge. The increase in tPA activity in the follicle base was transient, whereas the increased activity in the apex was maintained through the 24 h time point. Activity for uPA increased in the follicle apex and base within 12 h of the gonadotropin surge and remained elevated. Plasmin activity in follicular fluid also increased within 12 h after the preovulatory gonadotropin surge and was greatest at 24 h. Our results indicate that mRNA expression and enzyme activity for both tPA and uPA are increased in a temporally and spatially specific manner in bovine preovulatory follicles after exposure to a gonadotropin surge. Increased plasminogen activator and plasmin activity may be a contributing factor in the mechanisms of follicular rupture in cattle.

corpus luteum, follicle, granulosa cells, ovary, ovulation, theca cells


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The preovulatory gonadotropin surge initiates the ovulatory process and subsequent corpus luteum (CL) formation. Both of these events feature extracellular matrix (ECM) degradation and tissue remodeling. Such restructuring requires the targeted action of proteolytic enzymes. One such family of enzymes implicated in the preceding processes is the plasminogen activators. The plasminogen activator system includes plasminogen/plasmin, two specific plasminogen activators, cell surface plasminogen activator receptors, and several plasminogen activator inhibitors. Plasminogen is abundant in blood and peripheral tissues and, consequently, regulation of plasmin activity occurs at several levels. The plasminogen activators tissue plasminogen activator (tPA) and urokinase plasminogen activator (uPA) convert plasminogen into its active form, plasmin, in the extracellular milieu. Urokinase plasminogen activator can also bind to its cell surface receptor (uPAR) and form a stable complex for several hours [1, 2]. One function of the uPA-uPAR complex is to focus uPA-directed plasmin activity at the cell surface.

Degradation of the apical follicular layers is ultimately required for oocyte escape. Ovarian targets for plasmin include fibrin, fibrinogen, types III and IV collagen, fibronectin, laminin, and proteoglycans [3, 4]. In addition to playing a direct role in ECM degradation, plasmin can also activate the proenzyme form of several matrix metalloproteinases (MMPs) implicated in follicular rupture including MMP-1, MMP-3, and MMP-9 [5–9]. Furthermore, because of the relatively high concentration of plasminogen in tissue and body fluids, a small increase in plasminogen activator causes a large increase in plasmin in the extracellular milieu. Plasmin together with the MMPs can potentially degrade all ECM components in the ovary. Therefore, regulation of plasminogen activation may be a key regulatory step in the ovulatory process.

Several lines of evidence support a potential role of the plasminogen activators and plasmin in ovulation. Plasmin can decrease the tensile strength of the bovine follicle wall [10]. Furthermore, administration of antibodies against uPA [11] or tPA [12, 13] reduces the ovulation rate in sheep and rats, respectively. Before ovulation, tPA is the primary plasminogen activator induced in pig preovulatory follicles [14]. In contrast, uPA is the predominant plasminogen activator induced in mouse and sheep preovulatory follicles during the same time period [11, 15]. Thus, regulation of tPA and uPA in preovulatory follicles appears species-specific. Furthermore, the regulation and regulatory role of the plasminogen activator system during the periovulatory period in monotocous species, such as cattle, is not understood. Therefore, our objectives were to investigate the effect of the preovulatory gonadotropin surge on the localization and regulation of mRNAs for the plasminogen activators (tPA, uPA) and the cell surface receptor for uPA (uPAR) and on tPA, uPA, and plasmin activity in bovine preovulatory follicles. We observed a pronounced temporally and spatially specific increase in tPA, uPA, and uPAR mRNA levels and tPA, uPA, and plasmin activity in bovine periovulatory follicles in response to the gonadotropin surge. These results support a proposed role of gonadotropin surge-induced increases in both plasminogen activators in the ovulatory process and/or the morphologic changes associated with the ovulatory follicle-CL transition in cattle.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animal Care

Use of animals was approved by the All University Committee on Animal Use and Care at Michigan State University (Approval No. 04/98-056-00).

Experimental Model

Follicle development and timing of the preovulatory gonadotropin surge were synchronized in Holstein cows using the Ovsynch procedure (GnRH-7days-prostaglandinF2{alpha}[PGF2{alpha}]-36h-GnRH) [16]. Daily ultrasound analyses were performed after the first GnRH injection until the time of follicle collection to verify follicle synchrony and to exclude animals that turned over a new follicular wave before the second GnRH injection. Ovaries containing ovulatory follicles or new CL were collected by colpotomy at 0, 6, 12, 18, 24, and 48 h (follicles: 0, 6, 12, 18, and 24 h; 1-day-old CL = 48 h) after the second GnRH injection. Blood samples were collected at the time of PGF2{alpha} injection and at the time of the second GnRH injection. Serum progesterone concentrations in these samples were measured by RIA (Diagnostic Products Corporation, Los Angeles, CA) to ensure that all animals included in the study responded to the PGF2{alpha} injection with a decrease in serum progesterone, indicating CL regression. Intraassay and interassay coefficients of variation were 5.6% and 9.1%, respectively. To verify that none of the animals included in the study exhibited a preovulatory gonadotropin surge before the second GnRH injection, three blood samples were collected at 15-min intervals every 8 h beginning 16 h after the PGF2{alpha} injection and continuing until the time of ovariectomy or GnRH injection. A premature LH surge was not detected in any of the animals included in the 0 h (pre-gonadotropin surge) group. To confirm that a gonadotropin surge was elicited by the second GnRH injection, blood samples were also collected every hour for 4 h after the second GnRH injection. In the remaining animals, an LH surge occurred only after GnRH injection, verifying control of the timing of the gonadotropin surge in our model system. Concentrations of serum LH were measured by RIA [17, 18]. Intraassay and interassay coefficients of variation were 5.8% and 15.6%, respectively.

Tissue Collection

For mRNA quantification and enzyme activity assays, ovaries containing the ovulatory follicle or new CL were collected at 0, 6, 12, 18, 24, and 48 h (n = 5–6 at each time point; total 35) after the second GnRH injection. After ovariectomy, the ovulatory follicle or new CL was isolated by cutting away all remaining ovarian stroma and small follicles so that the ultrastructure at the apex of the follicle remained intact. Follicular fluid was aspirated, centrifuged, aliquoted, and stored at -20°C until activity assays. Follicles were then sagittally cut in half. One half was used for total RNA isolation. For protein analysis, the remaining half was cut transversely in two equal pieces, one containing the follicle apex and one containing the base. New CL collected 48 h after the GnRH injection were used only for mRNA analyses. Samples were frozen at -80°C within 15 min of ovariectomy. For in situ hybridization, ovaries containing the ovulatory follicles were collected at 0, 6, 12, and 24 h (n = 3 at each time point; total 12) after GnRH injection. Ovulatory follicles were dissected from the ovary, immediately immersed in embedding medium (Fisher Scientific, Chicago, IL), frozen over liquid nitrogen vapors, and stored at -80°C until sectioned.

Preparation of cDNA Probes for tPA, uPA, and uPAR

The cDNAs for tPA, uPA, uPAR, and ribosomal protein L19 (RPL19) (a housekeeping gene) have all been previously cloned in the bovine. Using the reported sequences (GenBank accession nos.: tPA, X85800; uPA, X85801; uPAR, S70635; RPL19, AF270675), oligonucleotide primers were prepared and used in combination with RNA isolated from bovine CL in the reverse transcription-polymerase chain reaction (RT-PCR) to amplify cDNAs that encode bovine tPA (328 base pairs [bp]), uPA (351 and 210 bp), uPAR (409 bp), and RPL19 (366 bp) The PCR products were subcloned into pBluescript SK(+) vectors (Stratagene, La Jolla, CA), and the identities and orientations of cDNAs in vectors were confirmed by fluorescent dye primer sequencing. Two separate cDNAs were generated for uPA for combined use for in situ hybridization analysis (described subsequently).

Characterization of tPA, uPA, and uPAR mRNA Abundance

Total RNA was isolated according to the manufacturer's instructions using the Trizol reagent (Invitrogen, Carlsbad, CA). To determine transcript size and number and to optimize specificity of hybridization conditions, approximately 15 µg of pooled RNA from each sample per time point was subjected to Northern analysis [19]. For quantitation of tPA, uPA, and uPAR mRNA abundance, 5 µg of total RNA from each sample was applied in duplicate to a Zeta probe nylon membrane (Bio-Rad, Hercules, CA) using a dot blot apparatus (Bio-Rad) [19]. Northern and dot blot analysis was then carried out using specific bovine tPA, uPA, uPAR, or RPL19 32P-labeled cDNA probes generated by the PCR. RPL19 was used for normalization purposes. Each 20-µl PCR reaction included 1x PCR buffer; 2.5 mM MgCl2; 1.6 µM each of dATP, dGTP, and dTTP (Invitrogen); 0.25 µM of each primer; 100 pg DNA template; 1.5 units Taq polymerase (Invitrogen); and 0.825 µM [32P]dCTP (3000 Ci/mmol; NEN Life Science Products, Boston, MA). The amplification conditions were 95°C for 5 min; 94°C for 0.5 min, 52°C for 1 min, 72°C for 1.5 min for 40 cycles; 72°C for 10 min; and hold at 4°C. After amplification, the PCR reactions were brought to 100 µl in NETS (150 mM sodium chloride, 10 mM EDTA, 50 mM Tris, 0.1% SDS), and the unincorporated 32P was removed by spun column chromatography through G-50 Sephadex minicolumns (Sigma, St. Louis, MO) [19]. The membranes were incubated overnight at 42°C in 25 ml of prehybridization buffer: 50% formamide, 5x saline-sodium citrate buffer (SSC) (single-strength is 0.15 M NaCl and 0.015 M sodium citrate, pH 7.0), 5x Denhardt solution (single-strength is 0.02% Ficoll, 0.02% polyvinylpyrrolidone, 0.02% BSA), 0.05 M sodium phosphate (pH 6.9), 0.1% SDS, and 250 µg/ml denatured herring sperm DNA. The prehybridization buffer was discarded, and 25 ml of fresh hybridization buffer (50% formamide, 5x SSC, 1x Denhardt solution, 0.02 M sodium phosphate, 0.1% SDS, 10% dextran sulfate, 100 µg/ml denatured herring sperm DNA, and 1 x 106 cpm labeled probe) was added, and the membranes were incubated overnight at 42°C. The membranes were then washed in wash solution 1 (1x SSC, 0.1% SDS, 0.1% sodium pyrophosphate) at 42°C for 15 min, followed by consecutive washes in wash solution 2 (0.1x SSC, 0.1% SDS, 0.1% sodium pyrophosphate) at 42°C and 47°C for 15 min each. After the washes, the filters were exposed to a phosphorimager cassette (Bio-Rad). After exposure (2–24 h), the cassette was scanned using a phosphorimager (Bio-Rad). After Northern analyses, the size of RNA transcripts was determined based on relative migration of RNA molecular weight markers (Roche, Indianapolis, IN). After hybridization for tPA, uPA, or uPAR, the membranes were then stripped and reprobed with the 32P-labeled RPL19 cDNA. Preliminary experiments demonstrated that RPL19 mRNA abundance in bovine preovulatory follicles and new CL was not regulated by the gonadotropin surge (P > 0.05, data not shown). Relative densitometric units for tPA, uPA, and uPAR were quantitated (from dot blots) and adjusted relative to RPL19 mRNA expression using Molecular Analyst version 1.5 software (Bio-Rad). Preliminary Northern blot experiments demonstrated that hybridization and washing conditions used in subsequent dot blot analyses were specific and yielded hybridization to single transcripts of the expected size for each mRNA of interest. Preliminary experiments also demonstrated that an increase in hybridization intensity was detected after hybridization of each cDNA to increasing amounts of sample RNA (1–10 µg).

In Situ Hybridization

Follicles were cut on a Leica cryostat (W. Nuhsbaum, McHenry, IL) into 12-µm sagittal sections and mounted onto positively charged slides (Fisher Scientific). A sagittal section allows a view of the cell types contained at both the apex and the base of the follicle. Before hybridization, sections were prewarmed to room temperature for 10 min, fixed in 3.7% formaldehyde in PBS for 5 min, rinsed twice in 2x SSC for 2 min each, incubated in 0.25% acetic anhydride in 0.1 M triethanolamine-HCl (pH 8.0) for 10 min, dehydrated in increasing concentrations of ethanol (70%, 80%, 95%, and 100%) for 2 min each, delipidated in absolute chloroform for 5 min, rinsed in 100% and 95% ethanol for 2 min each, and then air dried for 1 h. Hybridizations for each mRNA of interest were carried out on serial sections in triplicate using antisense and sense (negative controls) 35S- or 33P-labeled complementary RNA (cRNA) probes generated from previously described tPA, uPA, and uPAR cDNAs. Both antisense and sense [35S]UTP (1250 Ci/mmol, NEN Life Science Product; tPA) or [33P]UTP (3000 Ci/mmol; uPA and uPAR) cRNA probes were generated using linearized cDNA templates and an in vitro transcription kit (Stratagene) according to the manufacturer's directions. To increase sensitivity for uPA mRNA localization, two cRNA probes were generated from distinct uPA cDNAs and cohybridized to the same follicle sections. Both uPA cDNAs yielded identical results in Northern analysis (data not shown). The transcription reaction was incubated at 37°C for 1 h, and template DNA was removed by incubation with 20 units of RNase-free DNase (Stratagene) at 37°C for 15 min. After DNase treatment, the reaction was diluted to 100 µl with NETS, and unincorporated radionucleotides were removed as described previously. Before hybridization, labeled probes were diluted in hybridization buffer to a concentration of 1.0 x 106 cpm/ml. The hybridization buffer included 50% formamide, 0.3 M sodium chloride, 10 mM Tris (pH 8), 1 mM EDTA, 1x Denhardt solution, 50 mM dithiothreitol, 0.5 mg/ml yeast tRNA, and 10% dextran sulfate. Hybridizations were performed by adding 60 µl of diluted probe per slide and then incubating the slides in a humidified 55°C oven for 16 h. After hybridization, slides were washed twice by shaking in 2x SSC for 15 min at room temperature and treated with RNase A (50 µg/ml) in 2x SSC for 1 h at 37°C. Slides were then washed at 55°C in 2x SSC containing 0.1% ß-mercaptoethanol (ßME) for 15 min, 1x SSC/0.1% ßME for 15 min, 1x SSC/50% formamide/0.1% ßME for 30 min, and twice in 0.1x SSC/0.1% ßME for 15 min. The slides were then dehydrated in increasing ethanol concentrations (60%, 80%, 95%, and 100%), air-dried for 1 h, and then dipped in 50% NTB-2 emulsion (Eastman Kodak, Rochester, NY). Slides were exposed to autoradiographic emulsion for 10 (tPA) and 50 (uPA and uPAR) days at 4°C and then developed, followed by counterstaining with hematoxylin and eosin. The exposure time for detection of a given mRNA of interest was the same for all time points. Digital bright- and dark-field images were acquired on a Leica research microscope equipped with SPOT Model No. 1.1.0 camera and version 3.2.4 software (W. Nuhsbaum).

Enzyme Activity Assays

Follicle extracts were prepared using procedures previously described by Murdoch and McCormick [20]. Briefly, the apical or basal sections of follicles were homogenized using a polytron homogenizer (Fisher Scientific) in 800 µl of a solution containing 10 mM calcium chloride and 0.25% Triton X-100. The homogenates were then centrifuged at 9000 x g for 30 min at 4°C. The supernatants were collected and frozen at -20°C until assayed. The pellets were then resuspended in 200 µl of a solution containing 50 mM Tris, 0.15 M sodium chloride, and 0.1 M calcium chloride, pH 7.6, and heated at 60°C for 6 min. After heat treatment, the samples were centrifuged at 27 000 x g for 30 min at 4°C, and the supernatants were frozen at -20°C until assayed. Preliminary experiments demonstrated that minimal plasminogen activator activity was present in the second extract collected after heat treatment. Therefore, activity in these (heated) samples was not determined.

Casein zymography was conducted as described by Roche et al. [21] with slight modifications, to measure tPA, uPA, and plasmin activity in the follicle apex and base and follicular fluid. Plasminogen-free gels were used to confirm that the activity (bands) detected were plasminogen dependent. Bovine brain cerebellum (a rich source of tPA [22]) and ovarian surface epithelial cell conditioned media (a source of uPA [23]) were used as standards. Follicular fluid (1 µl) or follicle homogenates (10 µg protein; apex and base) were subjected to electrophoresis at 140 V for 115 min in 10% polyacrylamide gels containing 0.2% casein (Sigma), 0.1% SDS, and 25 mU/ml human plasminogen (Sigma). After electrophoresis, gels were washed once in 2.5% Triton X-100 for 45 min to remove SDS and then incubated in incubation buffer (50 mM Tris, 0.1 M sodium chloride pH 7.6) at 37°C for 17 h. The incubation buffer lacked the heavy metals required for MMP-dependent caseinolytic activity. The gels were then stained using 0.05% Coomassie blue in 10% acetic acid, 45% methanol for 2 h; briefly destained in 10% acetic acid, 45% methanol; and then fixed in 10% glycerol. Bands of activity were visualized as clear zones (where casein degradation occurred) across a dark background. Gels were photographed using a Gel Documentation System (Bio-Rad), and images were inverted for better visual contrast. For validation purposes, equal amounts of protein or volume of follicular fluid from individual samples were pooled within a time point, subjected to casein zymography, and photographed for depiction in appropriate figures. To derive individual estimates of tPA, uPA, and plasmin activity in each sample, all individual samples were run, and densitometric scanning was performed. Variation between gels was adjusted relative to differences in activity for tPA and uPA standards loaded on every gel. The intraassay (gel) and interassay (gel) coefficients of variation were 4% and 13%, respectively. Activity for tPA and uPA was not observed when plasminogen was omitted from gels. Addition of the specific uPA inhibitor, amiloride (1 mM; Sigma) to the incubation buffer attenuated bands of activity corresponding to uPA. Addition of the plasmin inhibitor aprotinin (2 µg/ml; Sigma) to the incubation buffer attenuated all plasminogen activator and plasmin activity (data not shown).

Statistical Analysis

Differences in mRNA abundance or enzyme activity were determined by one-way ANOVA using the General Linear Models procedure of Statistical Analysis Systems (version 8.0; SAS Institute Inc., Cary, NC). Individual comparisons of mean RNA abundance or enzyme activity were performed using the Fisher Protected Least Significant Differences test. When heterogeneity of variance was detected, data were log transformed before statistical analysis.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Regulation of tPA, uPA, and uPAR mRNA Abundance During the Periovulatory Period

Northern analyses detected single mRNA transcripts of 2.4, 2.2, and 1.3 kilobase (kb) for tPA, uPA, and uPAR, respectively (Fig. 1, A, C, and E). The relative abundance, as determined by dot blot analysis, of tPA, uPA, and uPAR mRNAs was increased in bovine preovulatory follicles after the gonadotropin surge, but the temporal regulation of each plasminogen activator system component was distinct. Messenger RNA for tPA increased within 6 h after the gonadotropin surge and remained elevated through the 24 h time point (P < 0.05). However, the increase in tPA mRNA was not maintained through the ovulatory follicle-CL transition (48 h; Fig. 1B). Relative abundance of uPA mRNA increased within 24 h after the gonadotropin surge and remained elevated in new CL (48 h; P < 0.05; Fig. 1D). Messenger RNA for uPAR was increased at 6, 12, 24, and 48 h relative to the 0 h time point (P < 0.05). Relative levels of uPAR mRNA were most dramatically increased near the time of ovulation and the follicular-luteal transition (15- and 32-fold increases at 24 and 48 h compared with 0 h, respectively; Fig. 1F).



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  		FIG. 1. Effect of a GnRH-induced gonadotropin surge on tPA, uPA, and uPAR mRNA abundance in bovine periovulatory follicular and luteal tissue. A) Northern analysis of tPA mRNA expression. Note hybridization to single 2.4-kb transcript. B) Effect of the preovulatory gonadotropin surge on relative levels of tPA mRNA in bovine preovulatory follicles and new CL. C) Northern analysis of uPA mRNA expression. Note hybridization to single 2.2-kb transcript. D) Effect of the preovulatory gonadotropin surge on relative levels of uPA mRNA in bovine preovulatory follicles and new CL. E) Northern analysis of uPAR mRNA expression. Note hybridization to a single 1.3-kb transcript. F) Effect of the preovulatory gonadotropin surge on relative levels of uPAR mRNA in bovine preovulatory follicles and new CL. Data (B, D, F) are expressed as relative units of mRNA per unit of RPL19 mRNA x 100 (n = 5–6 per time point; total 35). Because of the heterogeneity of variance, values for uPA mRNA (D) were log transformed before analysis. Data are expressed as the mean ± SEM (tPA, uPAR) and the mean ± average SEM (uPA). Time points without a common superscript are different at P < 0.05

Localization of tPA, uPA, and uPAR mRNAs in Bovine Preovulatory Follicles

The spatial regulation of tPA, uPA, and uPAR mRNA expression in response to the gonadotropin surge also was distinct. Messenger RNA for tPA was localized to the granulosal layer at all time points examined (Fig. 2, B, E, and H; 0, 6, and 24 h depicted). A low level of expression was also observed in the thecal layer of follicles collected near the time of ovulation (Fig. 2H; 24 h). Messenger RNA for uPA was observed in the granulosal and thecal layers at all time points examined (Fig. 3, B, E, and H; 0, 6, and 24 h depicted). Unlike the localization of tPA mRNA, the localization of uPA mRNA was heterogeneous, particularly in 24 h follicles, with additional prominent hybridization signals distributed throughout the thecal layer and the adjacent ovarian stroma (Fig. 3H). Messenger RNA for uPAR was localized primarily to both the granulosal and thecal cell layers of follicles collected at the 6 and 12 h time points (Fig. 4E; 6 h depicted), but with a lower level of expression also detected in the adjacent ovarian stroma of follicles collected at the 24 h time point (Fig. 4H). Localization of uPAR mRNA in the thecal layer and adjacent stroma of 24 h follicles (Fig. 4H) was heterogeneous and similar to that observed for uPA (Fig. 3H).



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  		FIG. 2. In situ localization of tPA mRNA within bovine periovulatory follicles collected at 0, 6, and 24 h after GnRH injection (magnification x120). Representative bright-field micrographs of preovulatory follicles collected at the 0 h (A), 6 h (D), and 24 h (G) time points and stained with hematoxylin and eosin. Representative dark-field micrographs of the corresponding bright-field sections of preovulatory follicles collected at the 0 h (B), 6 h (E), and 24 h (H) time points and hybridized with a 35S-labeled antisense tPA cRNA. Representative dark-field micrographs of corresponding adjacent serial sections of the same follicles collected at the 0 h (C), 6 h (F), and 24 h (I) time points and hybridized with a 35S-labeled sense tPA cRNA (n = 3 per time point; total 9). Note highest expression of tPA mRNA in granulosal layer, with additional localization in thecal layer of follicles collected at the 24 h time point



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  		FIG. 3. In situ localization of uPA mRNA within bovine periovulatory follicles collected at 0, 6, and 24 h after GnRH injection (magnification x120). Representative bright-field micrographs of preovulatory follicles collected at the 0 h (A), 6 h (D), and 24 h (G) time points and stained with hematoxylin and eosin. Representative dark-field micrographs of the corresponding bright-field sections of preovulatory follicles collected at the 0 h (B), 6 h (E), and 24 h (H) time points and hybridized with a 33P-labeled antisense uPA cRNA. Representative dark-field micrographs of corresponding adjacent serial sections of the same follicles collected at the 0 h (C), 6 h (F), and 24 h (I) time points and hybridized with a 33P-labeled sense uPA cRNA (n = 3 per time point; total 9). Note localization of uPA mRNA to both the granulosal and thecal layers of follicles collected at the 0 and 6 h time points, with additional heterogeneous expression in the thecal layer and adjacent ovarian stroma of follicles collected at the 24 h time point



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  		FIG. 4. In situ localization of uPAR mRNA within bovine periovulatory follicles collected at 0, 6, and 24 h after GnRH injection (magnification x120). Representative bright-field micrographs of preovulatory follicles collected at the 0 h (A), 6 h (D), and 24 h (G) time points and stained with hematoxylin and eosin. Representative dark-field micrographs of the corresponding bright-field sections of preovulatory follicles collected at the 0 h (B), 6 h (E), and 24 h (H) time points and hybridized with a 33P-labeled antisense uPAR cRNA. Representative dark-field micrographs of corresponding adjacent serial sections of the same follicles collected at the 0 h (C), 6 h (F), and 24 h (I) time points and hybridized with a 33P-labeled sense uPAR cRNA (n = 3 per time point; total 9). Note localization of uPAR mRNA primarily to the granulosal and thecal layers of follicles collected at the 6 h time point, with additional lower level of heterogeneous expression also detected in the adjacent ovarian stroma of follicles collected at the 24 h time point

Effect of the Gonadotropin Surge on tPA, uPA, and Plasmin Activity in Bovine Preovulatory Follicles

Activity for tPA in follicle homogenates was observed as a single band that comigrated with the tPA standard (Fig. 5, A and C). Activity for uPA was observed as a doublet (Fig. 5, A and C) that comigrated with the uPA standard, and plasmin activity was observed as multiple bands (Fig. 6A). The uPA doublet presumably corresponds to the single- and two-chain forms of uPA, as both bands of activity were inhibited by amiloride (data not shown). No low molecular weight uPA activity (30–35 Mr x 10-3) was observed in follicular fluid or homogenates. Plasmin activity was not detected in homogenates of the follicle apex or base even when 6-fold greater amounts of protein were loaded on gels (data not shown). Both tPA and uPA enzyme activity were increased within 12 h after the gonadotropin surge in both the follicle apex (Fig. 5B) and base (Fig. 5D). Activity for tPA was differentially regulated in the follicle apex versus the base. Activity for tPA remained elevated in the apex but decreased to presurge (0 h) levels in the base by the 24 h time point (Fig. 5D). Elevated uPA activity was maintained in both the apex and base through the 24 h time point (Fig. 5, B and D). In follicular fluid, tPA and plasmin activity were increased within 12 h after the gonadotropin surge and were also elevated at the 24 h time point (Fig. 6, A and B). Activity for uPA was not readily detectable in periovulatory follicular fluid (Fig. 6A).



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  		FIG. 5. Detection of tPA and uPA activity in bovine periovulatory follicles by casein zymography. A) Representative zymogram demonstrating tPA and uPA activity in the apex of bovine periovulatory follicles collected at 0, 6, 12, 18, and 24 h after GnRH injection (pooled samples, 10 µg protein per time point). B) Densitometric analysis of tPA and uPA activity (individual samples) in apex of bovine periovulatory follicles collected at 0, 6, 12, 18, and 24 h after GnRH injection. C) Representative zymogram demonstrating tPA and uPA activity in the base of bovine periovulatory follicles collected at 0, 6, 12, 18, and 24 h after GnRH injection (pooled samples, 10 µg protein per time point). D) Densitometric analysis of tPA and uPA activity (individual samples) in the base of bovine periovulatory follicles collected at 0, 6, 12, 18, and 24 h after GnRH injection. Note the single band of tPA activity and the doublet of uPA activity that comigrated with appropriate standards. Because of the heterogeneity of variance, the data were log transformed. Data are expressed as the mean ± average SEM (n = 5–6 per time point; total 27). Time points without a common superscript are different at P < 0.05.



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  		FIG. 6. Detection of tPA and plasmin activity in bovine periovulatory follicular fluid by casein zymography. A) Representative zymogram demonstrating tPA and plasmin activity in follicular fluid collected from bovine periovulatory follicles at 0, 6, 12, 18, and 24 h after GnRH injection (pooled samples, 1 µl follicular fluid per time point). B) Densitometric analysis of tPA and plasmin activity in follicular fluid of bovine periovulatory follicles collected at 0, 6, 12, 18, and 24 h after GnRH injection (individual samples). Note single band of tPA activity and multiple bands of plasmin activity. Because of the heterogeneity of variance, the data were log transformed. Data are expressed as the mean ± average SEM (n = 5–6 per time point; total 26). Time points without a common superscript are different at P < 0.05


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Although a potential role of the plasminogen activators in the ovulatory process has been well established, the species-specific and cell-specific regulation of individual components of the plasminogen activator system during the periovulatory period and their exact contribution to the ovulatory process are less well understood. Our results suggest that the regulation of plasminogen activator system components (tPA, uPA, and uPAR) during the periovulatory period in cattle is in many ways distinct from that reported to date in other species. In the present studies, mRNA for tPA, uPA, and uPAR in bovine preovulatory follicles was increased in response to the gonadotropin surge. In addition, tPA, uPA, and plasmin activity in preovulatory follicle homogenates and/or follicular fluid also was increased in response to the gonadotropin surge. Our results are consistent with a potential role of gonadotropin surge-induced up-regulation of the preceding plasminogen activator system components in mediating bovine follicle rupture and/or the morphologic changes associated with the ovulatory follicle-CL transition in cattle.

The observed prominent increase in mRNA abundance and enzyme activity for both tPA and uPA in bovine preovulatory follicles in response to the gonadotropin surge is in contrast to that observed in other species. In the mouse, sheep, and the pig, only one plasminogen activator or the other is markedly increased near the time of ovulation. In the mouse, uPA is the most abundant and dramatically up-regulated plasminogen activator after hCG injection [24]. In addition, plasminogen activator activity is reduced by 90% in the ovaries of uPA null mutant mice [25]. However, a small induction of tPA mRNA [25, 26] specifically in the thecal layer [15] has been reported in mice and is in fact sufficient to support normal ovulation in uPA-deficient mutant mice [25, 26]. In contrast, nearly all of the plasminogen activator activity in pig preovulatory follicles could be neutralized by tPA antibodies [27]. Results in sheep indicate that uPA is obligatory for ovulation, as intrafollicular injection of uPA antibodies, but not tPA antibodies, disrupt the ovulatory process [11]. In the rat, tPA mRNA expression is increased in response to the gonadotropin surge. However, the regulation of uPA expression during the periovulatory period in the rat is controversial. Li et al. [28] observed a decrease in uPA mRNA and protein levels in rat preovulatory follicles after exposure to hCG. In contrast, Macchione et al. [29] reported that both plasminogen activators are present in rat preovulatory follicles near the time of ovulation, but that the thecal and granulosal layers respond differently to the gonadotropin surge. In the above study [29], mRNA for tPA was increased in both the thecal and granulosal layers. However, uPA mRNA was increased in the thecal layer but decreased in the granulosal layer after exposure to the gonadotropin surge. In the present studies, tPA mRNA was localized primarily to the granulosal layer of bovine follicles, with only a very low level of expression detected in the thecal layer near the time of ovulation (within 24 h after the gonadotropin surge). In situ hybridization experiments did not reveal evidence of differential regulation of uPA mRNA in the granulosal and thecal layers of bovine follicles in response to the gonadotropin surge. Thus, although the temporal regulation of tPA and uPA mRNAs was clearly distinct, steady-state mRNA abundance for both plasminogen activators was clearly increased in bovine follicles after the gonadotropin surge.

Enzyme activity for both tPA and uPA was also increased in bovine follicles after exposure to the gonadotropin surge. The mechanisms that spatially regulate proteolysis of the preovulatory follicle wall and preferentially direct ECM degradation to the follicle apex are not clear. In the present study, spatial regulation of plasminogen activator activity by the preovulatory gonadotropin surge was examined by analysis of tPA and uPA activity within samples collected from the preovulatory follicle apex (the site of ovulation) versus the base. In the follicle apex, both tPA and uPA activity were increased and remained elevated through the 24 h time point. In the follicle base, uPA activity also remained elevated through the 24 h time point. In contrast, tPA activity in the follicle base peaked within 12 h after the gonadotropin surge but then decreased to presurge levels by 24 h. The mechanisms responsible for the differential regulation of tPA activity in the preovulatory follicle apex versus the base are not clear, but similar regional differences in tPA activity have also been observed in pig and rat preovulatory follicles [14, 30]. In contrast, Colgin and Murdoch [11] observed higher levels of uPA activity in the apex versus the base of preovulatory ovine follicles, and the ovarian surface epithelial cells were the primary source of the elevated uPA activity in the follicle apex [23]. The activity of uPA increased in bovine follicles after the gonadotropin surge, but unlike the differential spatial regulation noted for tPA, no disparity was evident between uPA activity in the follicle apex and that in the follicle base.

Levels of endogenous proteinase inhibitors dramatically influence activity of the plasminogen activators and plasmin in the extracellular milieu. Although our data on plasminogen activator inhibitor (PAI) activity do not support a role for PAI-1 or PAI-2 in the differential regulation of tPA activity in the follicle apex versus the base (unpublished data), further studies will be required to characterize gonadotropin surge-induced changes in the PAIs and proteinase inhibitors with an affinity for plasmin. Such information will provide a more complete understanding of the complex regulation of the plasminogen activator system during the periovulatory period and facilitate elucidation of the potential role of the individual components of the plasminogen activator system in follicle rupture and subsequent luteal formation.

One key role of the cell surface receptor for uPA (uPAR) is to localize pericellular plasmin activity. Our results indicate that uPAR mRNA abundance is increased in a cell-specific manner in response to the preovulatory gonadotropin surge. A transient increase in uPAR mRNA abundance was detected at the 6 and 12 h time points, and levels were subsequently increased again at 24 and 48 h. Messenger RNA for uPAR was localized primarily to the granulosal and thecal layers at the earlier time points (6 and 12 h), but a lower level of heterogeneous expression was also detected in the adjacent ovarian stroma of follicles collected at the 24 h time point. Similarly, during the periovulatory period in the rat, uPAR mRNA and protein are increased in both the granulosal cells and the residual ovarian tissue [28]. Interestingly, we observed a heterogeneous localization of both uPA and uPAR mRNAs within the thecal layer and adjacent ovarian stroma near the time of ovulation. Although the present experiments using in situ hybridization did not permit definitive identification of labeled cells, it will be of interest to further define the specific cell types in the thecal layer and adjacent stroma with intense expression of uPA and uPAR. Colocalization of uPA and uPAR has been observed previously in migrating endothelial cells [31] and infiltrating white blood cells [32]. Endothelial cell migration is a key component of luteal development, as capillaries must penetrate the avascular granulosal cell layer after ovulation and form the rich blood supply necessary to support luteal development [33]. Localization of uPA to endothelial cells near the site of capillary formation in developing CL has been reported previously [34].

We also detected an increase in plasmin activity in bovine follicular fluid collected after the gonadotropin surge. Multiple bands of plasmin activity (of similar Mr) have been observed in mouse ovarian homogenates [25] and human blood plasma [21]. The gonadotropin surge-induced increase in plasmin activity in follicular fluid can most likely be attributed to the observed increase in follicular fluid levels of tPA and enhanced activation of ubiquitous plasminogen in follicular fluid. Plasmin has been detected previously in the follicular fluid of cattle [35] and other species including the rabbit, horse, and pig [27, 36–38]. Plasmin in follicular fluid may help degrade high molecular weight proteoglycans, causing a decrease in follicular fluid viscosity that facilitates oocyte escape [39]. Plasmin-mediated degradation of fibrinogen [40–44] may also prevent premature blood clot formation in the follicular antrum before rupture. Liu et al. [45] proposed that plasmin may assist in cumulus expansion by terminating oocyte-cumulus cell communication. Thus, increased follicular fluid levels of plasmin may promote conditions that facilitate ovulatory release of the oocyte.

Plasmin may also play an important role in mediating the ECM degradation required for follicle rupture in cattle. In sheep, intrafollicular injection of the plasmin inhibitor {alpha}2-antiplasmin suppresses ovulation of preovulatory follicles [46]. A similar reduction of ovulation efficiency has also been observed with intrabursal injection of {alpha}2-antiplasmin in the rat [13]. Plasmin can directly degrade basement membrane ECM components including collagen IV, proteoglycans, laminin, and fibronectin [3, 4]. Interestingly, peak plasminogen-dependent plasmin activity has been detected in the stigma of rat preovulatory follicles 2 h before ovulation [47]. However, the plasmin-insensitive type I and III collagens of the thecal layer and tunica albuginea must also be degraded before ovulation. Here, plasmin may play a key role in activation of other ECM-degrading enzymes, such as interstitial collagenase (MMP-1) [48], that degrade type I and III collagen and may be crucial for ovulation. Messenger RNA for MMP-1 is increased in bovine preovulatory follicles after exposure to the gonadotropin surge [49], but a role for plasmin in activation of pro-MMP-1 during the periovulatory period in cattle remains to be established.

In summary, we have demonstrated that both plasminogen activators (tPA and uPA) as well as the cell surface receptor for uPA (uPAR) are up-regulated in bovine preovulatory follicles after the gonadotropin surge and in a temporally and spatially specific manner. These results support a potential role of tPA, uPA, and uPAR during the periovulatory period, although more investigation will be required to determine the requirement of the aforementioned plasminogen activator system components for ovulation and/or luteal formation in the bovine.


   ACKNOWLEDGMENTS
 
The authors wish to thank Isam Qahwash for his help with tissue collection. We are grateful to Theresa Doerr for her clerical assistance. The authors wish to thank Dr. Jim Ireland and Janet Ireland for reagents and assistance with the LH RIA.


   FOOTNOTES
 
First decision: 11 September 2001.

1 Supported by USDA 98-35203-6226 (G.W.S.) and the Michigan Agricultural Experiment Station. Back

2 Correspondence: George W. Smith, Michigan State University, Department of Animal Science, 1230D Anthony Hall, East Lansing, MI 48824-1225. FAX: 517 353 1699; smithge7{at}msu.edu Back

Accepted: December 4, 2001.

Received: August 17, 2001.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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