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a Institute of Physiology, Technical University of Munich, D-85350 Freising-Weihenstephan, Germany
b Department of Anatomy and Physiology, University of Hohenheim, D-70593 Stuttgart, Germany
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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corpus luteum, growth factors, ovary
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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Molecular cloning of the complementary DNAs has revealed that human VEGF may exist as one of four different molecular species of, respectively, 121, 165, 189, and 206 amino acids. Bovine VEGF is expected to be one amino acid shorter [8]. The VEGF isoforms are encoded by the same gene, through alternative splicing of mRNA. The resulting four polypeptides have strikingly different secretion patterns, which suggests multiple physiological roles for VEGF isoforms. The two smaller members of this family are secreted by cells and may act as paracrine, whereas the third and fourth are mostly cell associated and may act as autocrine, despite all members having an identical signal sequence [9]. The biological activities of VEGF are mediated through two high-affinity receptor tyrosine kinases [5]. The Flt-1 (fms-like-tyrosine kinase), or VEGFR-1, and the Flk-1 (fetal liver kinase-1), or VEGFR-2, possess seven immunoglobulin (Ig)-like domains in the extracellular domain, a single transmembrane region, and a consensus tyrosine kinase sequence that is interrupted by a kinase insert domain. Vascular endothelial growth factors bind to VEGFR-1 with a dissociation constant of approximately 10–20 pM [10]. However, VEGFR-2 has a somewhat lower affinity for VEGF; the Kd has been estimated to be approximately 75–125 pM [11]. The expression of VEGFR-1 and VEGFR-2 genes is largely restricted to the vascular endothelium [12, 13]. Monocytes, in contrast to the endothelium, express only the VEGFR-1. It is assumed that VEGFR-1 is the functional receptor for monocytes and endothelial cells [14]. In addition, VEGFR-1 is expressed in both proliferating and quiescent endothelial cells [15], suggesting a role in the maintenance of endothelial cells as well. Evidence for the presence of VEGF in the ovary has been reported for rats [13, 16], human [17], sheep [18], and cows [19–21].
To define further the underlying mechanisms regulating the dynamic vascular changes associated with the growth of CL and maintenance of CL function, we evaluated in detail the mRNA expression pattern of VEGF, its receptors, the content of VEGF in tissue, and the localization of VEGF in luteal tissue by collection of CL from different stages during the estrous cycle and pregnancy.
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MATERIAL AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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The CL from German Fleckvieh cows were collected at a local slaughterhouse within 10–20 min of slaughter. The stage of the estrous cycle was defined by macroscopic observation of the ovaries (follicles and CL) and the uterus (size, color, consistency, connective tissue, and mucus) [22]. The CL were accordingly assigned to the following stages: Days 1–2, Days 3–to 4, Days 5–7, Days 8–12, Days 13–18, Days 18 and later (after regression), and of pregnancy (the crown-rump length of the fetus was measured to evaluate the stage of pregnancy). Luteal tissue was frozen in liquid nitrogen or immersion-fixed immediately after collection. Tissue was kept frozen at -80°C for approximately 4 wk, until the CL were processed for molecular techniques or tissue extraction and RIA measurement.
Culture of Microvascular Endothelial Cells
Cytokeratin-negative endothelial cells, type 3, derived from the microvascular bed of the developing bovine CL were used and cultured as described elsewhere [23].
Tissue Extraction of VEGF
The CL (1 g wet weight) were transferred into 10 volumes of PBS containing 0.1% 1.4-dithiothreitol (Merck, Darmstadt, Germany) and homogenized in an ice bath by Ultra Turrax (Janke and Kunkel, Staufen, Germany). Five bursts of 15 sec at maximum speed, with 45-sec intervals of cooling between each burst, were applied. The homogenate was subsequently centrifuged at 2000 x g for 15 min at 4°C. The supernatant was separated in 1-ml aliquots and kept frozen until analysis.
Western Blot
The SDS-polyacrylamide gel electrophoreses were performed according to the method described by Laemmli [24]. The same antibody as used for the RIA and immunohistochemistry was utilized. For immunoblot analysis, the antiserum was diluted 1:10 000 and detected using goat anti-rabbit IgG-peroxidase conjugate (1:40 000) in combination with an enhanced chemiluminescence system (Amersham, UK).
Radioimmunoassay for VEGF
Concentrations of VEGF in homogenate supernatant were measured in 200-µl samples by RIA using a rabbit antiserum prepared in our laboratory and raised against recombinant bovine VEGF164 (supplied by D. Gospodarowicz, Chiron Corp., Berkeley, CA). The same antibody was also used for immunohistochemistry. The recombinant bovine VEGF164 (D. Gospodarowicz) was used for iodination by the iodogen method, as described elsewhere [11]. Labeled VEGF was separated from free iodine by a prepacked, disposable NAP-1O column containing Sephadex G-25 medium (Amersham-Pharmacia, Freiburg, Germany). Elution of the column was performed with 0.05 M phosphate buffer (pH 7.5) containing 0.2% sodium acid in 0.5-ml fractions into tubes containing 200 µl of 0.05 M phosphate buffer, 1% BSA, and 0.2% sodium acid. To fractions with labeled VEGF, glycerin was added to a final concentration of 50% and stored at -20°C until RIA. The tracer was stable for 3–4 mo. The incubation buffer for RIA was 3 M NaCl containing 1% BSA and 0.1% Triton-X-100 (pH 7.5). The antiserum was used at a final dilution of 1:400 000. Separation of bound and free VEGF was completed using the double-antibody technique and 6% polyethylene glycol 6000 (Serva, Heidelberg, Germany). The intra-assay variations were less than 6%, and the interassay variations were less than 14%. The ED50 of the assay was 0.6 ng/ml. Dilution of samples containing endogenous VEGF or added VEGF from blood plasma, cell culture medium, tissue extracts, or follicular fluid ran parallel to the standard curve. The average recovery of exogenous VEGF was 93% to 96%.
Fixation of CL Tissue for Immunohistochemistry
The CL were dissected into 0.5-cm-thick tissue slices, immersion-fixed in Bouin solution for 12 h, dehydrated in a graded series of ethanol, cleared in xylene, and embedded in paraffin wax using conventional procedures. Serial sections of 5-µm thickness were cut on a Leitz microtome (Leitz, Wetzlar, Germany) and processed for immunohistochemistry.
Immunohistochemistry
Immunohistochemical demonstration of VEGF was performed according to the ABC method as described by Hsu et al. [25]. To expose antigenic sites, dewaxed sections were heated four times to 95°C at 600 W in a microwave unit, maintained for 5 min, and allowed to cool for 20 min. Sections were then treated with hydrogen peroxide (1%) in methanol for 30 min to block endogenous peroxidase, normal goat serum diluted 1:10 in PBS for 20 min to reduce nonspecific background staining, polyclonal anti-VEGF antibody diluted 1:300 in PBS overnight at 4°C, biotinylated secondary antibody:goat anti-rabbit IgG (1:400 v/v) for 30 min at room temperature, StreptAB-HRP-complex (DAKO, Hamburg, Germany) for 30 min at room temperature, and then histochemical visualization of peroxidase using 3',3'-diaminobenzidine hydrochloride chromogen (Biotrend, Cologne, Germany) in 0.0006% hydrogen peroxide-0.05 M Tris buffer (pH 7.6) for 5 min in a dark room.
The specificity of the immunocytochemical reactions was assessed by several methods: 1) preadsorption test involving the respective antigen (VEGF165) at a concentration of 15 µg/ml in prediluted antiserum; 2) replacement of the primary antibody with buffer; 3) their substitution with normal goat IgG (dilution, 1:10); 4) incubation with diaminobenzidine reagent alone to exclude the possibility of nonsuppressed endogenous peroxidase activity. Elimination of specific staining of tissue elements in the controls demonstrated the specificity of the reactions.
RNA Isolation
Total RNA from CL tissue was isolated using the single-step method described by Chomczynski and Sacchi [26] utilizing TRIzolTM reagent (Gibco BRL, Rockville, MD). Total RNA from cultured endothelial cells was isolated using the NucleoSpin RNA kit (Macherey-Nagel, Dueren, Germany). The RNA was dissolved in water and spectroscopically quantified at 260 nm. Aliquots were subjected to 1% denaturing agarose gel electrophoresis and ethidium bromide staining to verify the quantity and quality of RNA.
Reverse Transcription-Polymerase Chain Reaction
Four micrograms of total RNA were used for RT-PCR to generate single-strand cDNA in a 60-µl reaction mixture as described elsewhere [27]. The optimal amount of total RNA for reverse transcription was evaluated by testing different RNA concentrations. The primers encoding the bovine sequences were used as described elsewhere (VEGF [15]; VEGFR-1, VEGFR-2, and Ubiquitin [28]) and commercially synthesized (Amersham-Pharmacia):
The VEGF primer permits detection of all four isoforms (186, 318, 390, and 441 bp) representing VEGF121, VEGF165, VEGF189, and VEGF206. The conditions for enzymatic amplification (RT-PCR) were established on a gradient cycler (Eppendorf, Hamburg, Germany). The VEGF, VEGFR-1, and VEGFR-2 PCR contained 10 mM Tris-HCl (pH 8.8), 50 mM KCl, 1.5 mM MgCl2, 0.1% Triton X-100, 0.6 µM of each primer, and 0.5 U of thermostable polymerase PrimeZyme (Biometra, Göttingen, Germany) to 3 µl of cDNA (final volume, 25 µl). Ubiquitin PCR was performed under the same conditions, but a higher concentration of primer (1.0 µM) was used. Samples for VEGF, VEGFR-1, and VEGFR-2 were amplified for 35, 30, or 28 cycles, respectively, involving an initial denaturation step at 94°C for 2 min and then each cycle at 94°C for 1 min, 60°C for 1 min (VEGF at 55°C), and afterward, one additional elongation step at 72°C for 2 min. Samples for the housekeeping gene ubiquitin were amplified for 22 cycles, involving a single denaturation step at 94°C for 2 min and then each cycle at 94°C for 45 sec, 55°C for 45 sec, and afterward, one additional elongation step at 72°C for 2 min.
To determine the optimal quantity of reverse transcript needed for PCR and to verify that the cDNA product was dependent on the input of transcript, varying quantities of transcript were used in the PCR reaction. The reverse transcription product from 3 µl was in the linear range for this amount and produced a visible band. To exclude any contaminating genomic DNA, all experiments included controls lacking the reverse transcription enzyme. As a negative control, water was used instead of RNA for the RT-PCR to exclude any contamination from buffers and tubes.
Aliquots (5 µl) of the PCR reaction products were fractionated by electrophoresis through a 1.5% agarose gel containing ethidium bromide in a constant 60-V field. To determine the length of the products, a Mass Ladder and 100-bp marker (Gibco BRL) was used. The ethidium bromide-stained gels were evaluated by a video documentation system (Amersham-Pharmacia). For comparison of treatment effects, all gels being compared were run and stained at the same time to ensure that any apparent differences detected were, indeed, due to treatments and not between-run variability. Band intensities (relative) were analyzed by computerized densitometry using the Image Master program (Amersham-Pharmacia). Confirmation of the PCR product identity was obtained by cDNA subcloning into a transcription vector (pCR-Script; Stratagene, La Jolla, CA) and then subjecting them to commercial DNA sequencing (TopLab, Munich, Germany).
Ribonuclease Protection Assay
Total RNA (30 µg) was introduced into a commercial ribonuclease protection assay (RPA) II kit (Ambion, Austin, TX) and both performed and validated according to the method described by Gabler et al. [28]. Transcripts generated from linearized gene-containing plasmids by RNA polymerases (Stratagene) generated antisense RNA probes labeled with a 32P-cytidine triphosphate (CTP, 800 Ci/mmol, Amersham; RNA transcription kit, Stratagene). Unincorporated radioactive CTP was removed by centrifugation through MicroSpin columns (Amersham-Pharmacia). After hybridization (20 h), RNase digestion buffer (2.5 U/ml RNase A and 100 U/ml RNase T1) was added to each sample. Protected mRNA fragments were identified by horizontal polyacrylamide gel electrophoresis as described elsewhere [27]. After drying of the gels, autoradiography was performed using ECL films (Amersham-Pharmacia) for 5 days, 14 days, and 30 days. Band intensities were analyzed by computerized densitometry using the Image Master program (Amersham-Pharmacia). To determine the mRNA contents of the investigated factor, increasing quantities of in vitro synthesized and standardized sense-RNA were introduced in each RPA. Sense cRNA transcripts were generated from the cloned inserts using the appropriate T3 or T7 RNA polymerase (Stratagene). The RNA concentrations were estimated by comparing signal intensities with known standards.
Statistical Analyses
The statistical significance of differences in mRNA expressions and protein concentration of examined factors were assessed by ANOVA followed by Fisher least significant difference as a multiple-comparison test. All experimental data are shown as the mean ± SEM.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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Introduction experiments verified the specific transcripts for VEGF, VEGFR-1, and VEGFR-2 present in bovine CL and cultured endothelial cells by RT-PCR (data not shown). Each PCR product showed 100% homology to the known bovine genes after sequencing. To confirm the integrity of the mRNA templates and RT-PCR protocol, the housekeeping gene ubiquitin was examined in all samples. A representative example for the ubiquitin-specific RT-PCR products (189 + 417 bp) is shown in Figure 1A. The relative signal intensities for PCR products specific for VEGF, VEGFR-1, and VEGFR-2 were assessed after correction based on the ubiquitin PCR signal intensities.
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Analysis of the VEGF transcripts by RT-PCR showed that CL predominantly expressed the two smallest VEGF isoforms (VEGF121 and VEGF165) and very weakly expressed the VEGF189 isoform, as shown in Figure 1A. The results of the densitometric analysis of mRNA expression by RT-PCR for VEGF examined in CL are shown in Figure 1B. The highest VEGF mRNA expression was detected during the early luteal phase of the estrous cycle (period of angiogenesis) and during pregnancy. The expression intensity of VEGF decreased significantly after Day 7 to lowest level after regression of CL. Concerning the target cells for VEGF expression, RNA obtained from CL tissue and cultured CL endothelial cells was also investigated. Figure 1C shows a representative example for VEGF RT-PCR expression in CL tissue (strong signal) compared with endothelial cells (weak signal), suggesting that luteal cells may be the main source of VEGF mRNA production. The VEGFR-1 mRNA expression data by RT-PCR and RPA are presented in Figure 2, showing no obvious differences in expression during the different phases examined. In contrast, the RT-PCR expression pattern for VEGFR-2 (Fig. 3A) shows a clear regulation, with the highest level occurring during early luteal phase (Days 3–4) and then a significant decrease afterward, with a slight increase during pregnancy. This pattern of VEGFR-2 mRNA expression was further confirmed by RPA (Fig. 3B).
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VEGF RIA and Protein Tissue Concentration
Data for characterization of the antibody by Western blotting or RIA are shown in Figure 4. Western blotting (Fig. 4A) shows two bands for recombinant human (rh) VEGF165 and recombinant bovine (rb) VEGF164 (glycosylated and unglycosylated) and one band for rhVEGF189. As shown in lane 5, the two bands for rbVEGF164 disappeared after preadsorbtion of the diluted antibody with 15 µg of rhVEGF165 per milliliter.
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A typical standard curve for VEGF RIA is shown in Figure 4B. A strong cross-reaction with rhVEGF165 was observed. The cross-reactivity to other growth factors, such as platelet-derived growth factors (PDGF), rhPDGF-AA, rhPDGF-BB, rhPDGF-AB, and recombinant human transforming growth factor-
(Pepro Tech, Inc., Rocky Hill, NJ), or rbFGF1 and rbFGF2 (D. Gospodarowicz), were less than 0.1%.
The VEGF protein concentration measured in CL extracts without heparin sepharose column separation by RIA is presented in Figure 5. The highest levels were measured during the early luteal phase, followed by a significant decrease afterward (especially after regression). The CL of pregnancy still contained clear, measurable (but lower) levels compared with the cyclic functional CL.
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Immunohistochemistry
After ovulation (Days 1–2), VEGF immunoreactivity was seen in a subset of luteal cells and partially in blood vessels (Fig. 6A). Figure 6, B (Days 5–7) as well as C and D (Days 13–16), shows VEGF in luteal cells with strong differences in the intensity of immunostaining between adjacent luteal cells. As demonstrated in Figure 6D (high-power micrograph), the VEGF protein can be localized in small granules in a subset of strongly labeled luteal cells. In contrast, in regressing luteal tissue (Fig. 5, E and F), intensive VEGF immunoreaction is restricted to the cytoplasm of the smooth muscle cells of blood vessels. Endothelial cells as well as luteal cells were consistently negative at this cycle stage.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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The mRNA expression for VEGF in the bovine CL agrees with observations during in situ hybridization of rat luteal cells [13] or in primate CL during the early and late luteal phase by Northern analysis or in situ hybridization [31]. The highest mRNA expression levels were observed, as evaluated by RPA, in sheep CL on Days 2–4, on Day 8, or on Days 14–15 of the estrous cycle [18], which agrees with our VEGF mRNA expression data and protein tissue concentrations in the cow. A higher mRNA expression was also found during the early luteal phase in bovine CL by RT-PCR [21]. The localization of VEGF in luteal cells was also observed in human and bovine CL [21, 32]. However, in contrast to the bovine CL, immunostaining in human granulosa and theca lutein cells became weak in the mid and late luteal phase [32]. We could show in the regressed CL that mRNA expression and tissue levels decreased significantly. No hybridization signals were observed from regressing CL in primates [31]. Immunostaining of luteal cells from regressing CL disappeared in our study, which agrees with other observations in the bovine CL [21]. However, in contrast, we could observe some positive staining for VEGF in the smooth muscle cells of arteries (Fig. 6, E and F).
At the moment, we do not have a clear explanation for this phenomenon, but hypoxia could explain this expression [33]. Concerning regulation of VEGF expression and protein production, oxygen tension plays a major role, both in vitro and in vivo. The VEGF mRNA expression is rapidly and reversible induced by exposure to low PO2 in a variety of normal and transformed cultured cell types [33–36], and LH is also a potent stimulator of VEGF mRNA expression in bovine granulosa cells [19]. Recently, we demonstrated the stimulation of VEGF secretion in bovine granulosa cells by LH and insulin-like growth factor I, hormones that are known to be most effective for luteinization [37]. The gonadotropic regulation of the expression of mRNA for VEGF in the CL was studied in vivo by treating monkeys with a potent GnRH antagonist during the mid luteal phase of the menstrual cycle. A 3-day treatment regimen brought about significant reduction in the levels of mRNA for VEGF [31]. In rats treated with hCG, increases of VEGF transcripts were detectable both in granulosa cells and in thecal/stromal tissue and the early CL [16]. Cell differentiation itself plays an important role in the regulation of VEGF gene expression [38]. This could be an additional stimuli for VEGF expression in vivo during the transformation of granulosa and theca cells to luteal cells. The same signaling pathway by stimulation of cAMP production and protein kinase A activation for progesterone secretion (luteinization) and VEGF secretion (angiogenesis) demonstrates a close, temporal relationship of these normal physiological processes. Angiogenesis, the formation of new capillaries (sprouting from pre-existing ones), is one of the prominent features of the early CL, and VEGF is the most important factor in the regulation of both normal and abnormal angiogenesis [5]. Treatment with truncated soluble VEGFR-1, which inhibits VEGF bioactivity, resulted in virtually complete suppression of CL angiogenesis in a rat model of hormonally induced ovulation; this effect was associated with inhibition of CL development and progesterone release [39].
Growth factor activation enables quiescent, resting endothelial cells to proteolytically degrade their underlying extracellular matrix, to invade and directionally migrate toward the angiogenic stimulus, and to proliferate and organize into new, three-dimensional capillaries [40]. Our finding that VEGF is predominantly localized in luteal cells underlines these assumptions, suggesting that VEGF may act as a chemoattractant for sprouting endothelial cells. This idea is supported by the dominant expression of the secretory forms of VEGF121 and VEGF165 (Fig. 1A).
As in our study, rhVEGF120, rhVEGF164, and rhVEGF188 isoforms have been found in sheep CL [18]. Evaluation from the RPA showed that VEGF120 represented approximately one-third of the total mRNA encoding VEGF in the CL, and that this proportion did not vary with the stage of the estrous cycle [18]. The targets for VEGF localized in luteal cells are endothelial cells, where both receptors (VEGFR-1 and VEGFR-2) are found. Probably, the growth-stimulatory signals are transducted, to a major extent, via VEGFR-2. This type of receptor is clearly regulated in the bovine CL, despite our results that present a high expression of VEGFR-1. Because both receptors bind VEGF with high affinity [10, 11], coexpression of VEGFR-1 and VEGFR-2 in endothelial cells may lead to formation of heterodimers in response to VEGF. Porcine aortic endothelial cells, individually expressing VEGFR-1 or VEGFR-2 after transfection, differed in their ability to migrate toward VEGF [41]. The VEGFR-1-expressing cells failed to migrate, whereas the VEGFR-2-expressing cells migrated efficiently, with a maximal response at 10 ng/ml VEGF. Thus, it is likely that VEGFR-2 expressed in endothelial cells could mediate chemotaxis.
In the present study, we demonstrate that VEGF and its receptors are clearly expressed in the bovine CL during the estrous cycle and pregnancy. The highest mRNA expression for VEGF and VEGFR-2 was detected during the early luteal phase, correlating with the highest VEGF protein tissue concentrations and localization in luteal cells and coinciding with luteal vascularization. The most potent vascular permeability agent, VEGF possesses 50 000-fold greater potency than histamine [42], with an effective concentration below 1 nM. Microvessels that supply the CL are reported to be among the leakiest in the body, possibly due to the action of VEGF action [43, 44]. The close temporal and spatial correlation between VEGF expression by luteal cells and local-vessel hyperpermeability suggests that VEGF is the responsible mediator. This specific effect may possibly be mediated by VEGFR-2.
For the formation of new vessels, the basement membrane must be degraded to allow migration of endothelial cells into the surrounding tissue. Vascular endothelial growth factor has been shown to induce plasminogen activators [45]. In the porcine model [41], VEGF stimulate urikinase plasminogen activator production in VEGFR-1-expressing endothelial cells but not in VEGFR-2-expressing cells [46]. Angiogenesis in the newly formed bovine CL is normally completed at Days 5–7 of the estrous cycle. The relative high expression of VEGF and its receptors afterward and in the CL of pregnancy (nonangiogenic phase) suggests a maintenance function of VEGF for the endothelial cells of the surrounding capillaries or luteal cells themselves. As shown by Alan et al. [47], a certain threshold concentration of VEGF is required to inhibit apoptosis of the endothelial cells and is essential for stabilization of the newly formed blood vessels.
Basic FGF (FGF2) is another important factor for angiogenesis. Under in vitro conditions and compared with FGF2 at equimolar concentrations, VEGF was approximately half as potent. However, the most striking effect was seen in combination with FGF2: When added simultaneously, VEGF and FGF2 induced an in vitro angiogenic response that was far greater than additive and that occurred with greater rapidity than the response to either cytokine alone [48]. Those authors suggest that the synergism between VEGF and FGF2 plays an important role in the control of angiogenesis in vivo. In the early bovine CL (Days 2–3), intense immunostaining for FGF2 is exclusively observed in vascular cells; luteal cells are negative [49]. With increasing age of the CL, the staining pattern for FGF2 changes dramatically. On Days 4–7 (end of angiogenesis), the large luteal cells exhibit a very intense and distinct labeling of the cell membrane, and the vascular cells still stain positive. Beginning with the mid luteal phase (Days 8–12), most of the capillary endothelial cells are no longer positive for FGF2, and FGF2 is now localized exclusively in the cytoplasm of the luteal cells. These results underline the importance of the in vitro data and participation of FGF2 and VEGF for angiogenesis.
In conclusion, the results presented of mRNA expression for VEGF, its isoforms and receptors, and VEGF protein tissue concentration and localization suggest an important role of the VEGF system in conjunction with FGF2 in angiogenesis of the newly forming CL. The maintenance of capillary function surrounding the luteal cells could be regulated through this growth factor system, again together with FGF2 by autocrine/paracrine mechanisms in the bovine CL during the estrous cycle and pregnancy.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by the German Research Foundation (Scha 257/14-1). 
2 Correspondence: Dieter Schams, Institute of Physiology, Technical University of Munich, Weihenstephaner Berg 3, D-85350 Freising-Weihenstephan, Germany. FAX: 49 8161 714204;physio{at}weihenstephan.de
Accepted: May 18, 2000.
Received: December 28, 1999.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIAL AND METHODSRESULTSDISCUSSIONREFERENCES |
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H. Cardenas, K.A. Burke, R.M. Bigsby, W.F. Pope, and K.P. Nephew Estrogen Receptor {beta} in the Sheep Ovary During the Estrous Cycle and Early Pregnancy Biol Reprod, July 1, 2001; 65(1): 128 - 134. [Abstract] [Full Text] [PDF]
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