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Biology of Reproduction 65, 1672-1680 (2001)
© 2001 Society for the Study of Reproduction, Inc.

Regular Article

Localization of the Renin-Angiotensin System in the Bovine Ovary: Cyclic Variation of the Angiotensin II Receptor Expression1

K.H. Schauser2,a, A.H. Nielsena, H. Winthera, V. Dantzera, and K. Poulsena

a Department of Anatomy and Physiology, The Royal Veterinary and Agricultural University, DK-1870 Frederiksberg C, Denmark


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Angiotensin (Ang) II may modulate reproductive function in the bovine ovary. Therefore, expression and localization of a local ovarian renin-angiotensin system (RAS) were investigated by elucidating the influence of the estrus cycle, pregnancy, and the presence of follicular cysts. Receptor analysis and autoradiography were used to characterize and localize Ang II receptors. Cyclic variations in the density of ovarian Ang II receptors were found with a higher value in estrus than in diestrus. The density in ovaries with follicular cysts was in the same order of magnitude as in estrus. The Ang II receptor type 2 (AT2) dominated in all three groups. Autoradiography showed that the majority of antral follicles and follicular cysts had intense AT2 receptor binding in the theca externa. Binding was less intense in the theca interna, whereas there was no binding in the granulosa layer. In the corpora lutea, the AT2 receptor was dominant in the capsule and in connective tissue infoldings, whereas no binding was observed in the luteal tissue. The type 1 Ang II receptor (AT1) was dominant in the stroma and showed no cyclic changes. Angiotensin-converting enzyme (ACE) activity was detected in all aspirated follicular fluids and homogenates of ovarian tissue. Autoradiography showed that most of the ACE was localized on endothelial cells. Renin immunoreactivity was found in granulosa and thecal cells of antral follicles and in luteal cells. Furthermore, solitary cells in the stroma, presumably macrophages, displayed intense staining. Our finding of cyclic changes support the concept of an active and regulated RAS in the bovine ovary.

corpus luteum, follicle, ovary, ovulatory cycle, theca cells


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The renin-angiotensin system (RAS) is well known for its systemic role in the regulation of blood pressure and fluid homeostasis. Recently, several organs were shown to contain a local RAS [1]. In the ovary of various mammalian species such as rabbit, rat, and mouse, all components of an RAS were identified, indicating the existence of a local ovarian RAS [2, 3].

As recently as 1988, Lightman et al. [4] found cycle-related changes of angiotensin (Ang) II receptor expression in rat ovaries with the highest expression in diestrus. The two major types of Ang II receptors have been designated AT1 and AT2 [5]. In rat ovaries, the AT1 receptor was primarily found in the stroma, whereas AT2 was expressed mostly on granulosa cells of atretic follicles [6]. In contrast, rabbit granulosa cells express high levels of AT2 receptors in the preovulatory follicle [7]. In bovine ovarian follicles, Ang II receptor expression—mainly type AT2—correlated positively with follicular diameter and tissue weight [8]. In another study of bovine granulosa, thecal, and luteal cells, Ang II binding sites were found exclusively in thecal cells [9]. This difference in Ang II receptor localization prompted an investigation of the species specificity of the ovarian RAS because the system components show a variation in their distribution and concentration among species [10, 11].

The possibility that locally produced Ang II serves as a significant autocrine and paracrine modulator in oocyte maturation and the process of ovulation has been discussed before. Pellicer et al. in 1988 [12] demonstrated that Ang II was involved in ovulation in the rat, whereas Daud et al. [13] found no Ang II influence on this process. There is evidence that Ang II modulates steroidogenesis in the rat ovary [14, 15]. It has been suggested that in both rat and rabbit, the AT2 receptor induces granulosa cell apoptosis and follicular atresia [16, 17]. The effect of Ang II in the bovine ovary is still unknown, but in an in vitro study with bovine luteal cells, Ang II reduced LH-induced progesterone production and had a stimulatory effect on mRNA expression of basic fibroblast growth factor (bFGF) [18]. Microdialysis of bovine preovulatory follicles demonstrated a complex interaction between Ang II and other vasoactive peptides [19], and in bovine mature follicles Ang II infusion stimulated the release of progesterone, estradiol, and prostaglandins [20]. These findings suggest that Ang II in the bovine ovary may play important regulatory roles in reproductive function by affecting oocyte maturation, ovulation, and the expression of reproductive hormones and vasoactive substances.

In order to study the local RAS in the bovine ovary and to elucidate changes during the estrus cycle and pregnancy, and the effect of the presence of follicular cysts, we determined the distribution and expression of the two Ang-forming enzymes, renin and angiotensin-converting enzyme (ACE), and the Ang II receptors. This was accomplished using 1) cell membrane fractions of ovarian tissue to determine the characteristics and density of Ang II receptors and ACE activity, 2) autoradiography to localize and characterize the Ang II receptor types and to localize ACE, and 3) immunohistochemistry to localize renin.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Collection of Tissues

Bovine ovaries were collected at an abattoir 20–30 min after exsanguination. The ovaries of animals that demonstrated an apparently normal cycle were classified by gross appearance on the basis of viscosity and amount of cervical and vaginal mucus as described earlier [21]. Ovaries from pregnant cows were collected and the gestational stage was estimated by the fetal crown-rump length [22]. Ovaries with follicular cysts were defined as having more than two cysts larger than 20 mm.

Tissue samples for Ang II receptor binding studies were transported to the laboratory on ice (1–2 h) and stored at -20°C. Tissue samples for measurement of ACE activity were transported on dry ice and stored at -80°C. For analysis of whole ovaries, the entire ovary was stored frozen, thawed just before use, and cut in pieces at 0°C. Tissue pieces used for homogenization were randomly collected. For measurement of ACE activity on pools of antral follicles, all obtainable ovaries with no pathological changes were collected without selection and brought to the laboratory (1–2 h at 37°C). All follicles (>4 mm) from that day were dissected on ice, pooled, and stored at -80°C. Follicular fluids were immediately aspirated and frozen in liquid nitrogen. For autoradiography, tissue samples (1–3 mm) were rapidly removed, snap-frozen in liquid nitrogen, wrapped in Parafilm (American Can Company, Greenwich, CT), transported on dry ice, and stored at -80°C. For immunohistochemistry, tissue samples (2–3 mm) were rapidly removed and immediately fixed in Bouin solution (Bie & Berntsen, Rødovre, Denmark) containing 10.25% formaldehyde, 4.81% acetic acid, and 0.88% picric acid for 18–24 h at 4°C. After dehydration through a graded series of alcohol and xylene or estisol 220 (Esti Chem, Køge, Denmark), tissues were embedded in paraffin wax.

Angiotensin II Receptor Analysis in Cell Membrane Fractions

Cell membrane fractions for Ang II receptor binding studies were prepared as previously described in detail [21, 23]. The membrane fractions were frozen at -20°C for later analysis.

The Ang II receptor assay was performed in a final volume of 150 µl by incubation at 37°C for 1 h as described earlier [21, 23]. The resuspended membrane fraction (100 µl) was incubated with 25 µl of 125I-[Sar1-Ile5-Ile8]-Ang II (specific activity, 2200 Ci/mmol; DuPont, NEN Research Products, Wilmington, DE) in a buffer containing 0.5 g/L soybean trypsin inhibitor, type I-S (Sigma, St. Louis, MO), 0.6 g/L bacitracin (Sigma), and 2 g/L human serum albumin (Statens Serum Institut, Copenhagen, Denmark). The density of Ang II binding sites and their binding characteristics for [Sar1-Ile5-Ile8]-Ang II were determined by using 25 µl of varying concentrations of unlabeled [Sar1-Ile5-Ile8]-Ang II (Sigma) as a ligand. For characterization of the Ang II receptor types [24, 25], 25 µl of varying concentrations of the type-specific Ang II receptor antagonist, losartan (DuP 753; Du Pont Merck Pharmaceutical Company, Wilmington, DE) or PD123319 (Parke-Davis, Ann Arbor, MI), were used as ligands as described before [23]. After incubation for 1 h, 3 ml of ice-cold 10 mM sodium phosphate (pH 7.4) containing 120 mM sodium chloride was added, and the mixture was immediately filtered through Whatman GF/F glass fiber filters (Whatman, Clifton, NJ) presoaked with 2 g/L human serum albumin. The filters were washed twice with 3 ml of the same buffer and counted in a gamma counter. The KELL collection of programs (Radlig version 6.0.5; Biosoft, Cambridge, U.K.) was used to analyze the radioligand binding experiments. The apparent dissociation constant (Kd) and the maximal binding capacity of ligand (Bmax) were calculated by Scatchard analysis. The Ang II receptor density was calculated by using Bmax and the protein concentration.

Angiotensin II Receptor Autoradiography

An in situ autoradiographic technique as described earlier in detail by Schauser et al. [23] was used. Briefly, frozen sections (10 µm) were brought to room temperature and incubated for 3 h in a buffer containing 0.5 nM 125I-[Sar1-Ile5-Ile8]-Ang II (DuPont, NEN Research Products). Nonspecific binding was determined on parallel sections using identical incubation conditions except for the addition of 1 µM unlabeled [Sar1-Ile5-Ile8]-Ang II. For localization and characterization of the Ang II receptor types, parallel sections were incubated in the same buffer containing 0.5 nM 125I-[Sar1-Ile5-Ile8]-Ang II in the presence of 5 µM losartan, 5 µM PD123319, or both. The sections were dipped in a K2 photographic emulsion (Ilford, Brønshøj, Denmark). The Ang II receptor density was estimated by a visual subjective scoring of the deposition of silver grains in the autoradiograms.

Measurement of ACE Activity

A protocol as described by Cushman et al. [26] was used. Analysis of the ACE activity in follicular fluid was performed directly on the aspirated fluid, diluted 1:10 in 50 mM potassium phosphate (pH 7.8). For analysis on ovarian tissue, 2 g of the tissue was homogenized at 0°C in 8 ml of 50 mM potassium phosphate (pH 7.8) for 30 sec with an Ultra-Turrax (IKA Labortechnik, Staufen, Germany) and then for 1 min at 200 x g with a Potter-Elvehjem homogenizer. The homogenate was centrifuged at 700 x g for 5 min at 4°C and the supernatant obtained was stored at -80°C for later analysis.

All measurements were made in duplicate. In this assay, hippuryl-l-histidyl-l-leucine (Sigma) is cleaved by ACE during incubation at 37°C. The product, histidyl-leucine was made fluorescent by reacting it with o-phthaldialdehyde (Sigma) and measured in a spectrofluorophotometer (RF-5001PC, Shimadzu, SpectraChrom, Brønshøj, Denmark) at 365 nm excitation and 486 nm emission wave lengths. The specificity of the ACE activity was verified by inhibition with the ACE inhibitors lisinopril (Sigma) or captopril (Sigma).

Autoradiographic Localization of ACE

Frozen sections (10 µm) were stored at -80°C up to 3 wk. Two different protocols, as described by Mow et al. [27] and by Krebs et al. [28], were used.

According to the first protocol, the sections were preincubated for 15 min in a buffer containing 10 mM sodium phosphate (pH 7.4) and 150 mM sodium chloride, and then incubated in a humidified chamber for 60 min at room temperature in a fresh buffer containing 10 mM sodium phosphate (pH 7.4), 150 mM sodium chloride, 2 g/L proteinase-free bovine serum albumin (Biofac A/S), and 0.5 nM 125I-lisinopril (Sigma, custom labeled by Nycomed Amersham plc, Buckinghamshire, U.K.). Nonspecific binding was determined on an adjacent section using identical incubation conditions except for the addition of 1 µM of unlabeled lisinopril (Sigma). After incubation, the sections were rinsed four times for 1 min each in ice-cold 50 mM Tris-HCl (pH 7.4) followed by a dip in ice-cold distilled water.

According to the second protocol, the sections were preincubated for 30 min at 22°C in a buffer containing 50 mM Hepes (Sigma), 150 mM NaCl, 10 µM ZnS04 pH 7.5, and incubated for 1 h in an identical buffer containing 0.3 nM 125I-MK-351A (Peptide Radioiodination Service Center, Washington State University, Pullman, WA). Nonspecific binding was determined by adding 1 µM unlabeled captopril (Sigma). After incubation, the sections were washed three times for 2 min each in ice-cold isotonic 50 mM Tris-HCl (pH 7.4) followed by a dip in ice-cold distilled water. All sections were quickly dried under a stream of hot air. Then they were either placed in a Hypercassette (Nycomed Amersham plc) and apposed to a Hyperfilm-3H (Amersham Denmark ApS, Birkerød, Denmark), or fixed and dipped in liquid emulsion Ilford K2. Rat small intestine served as a positive control. The ACE density was estimated by a visual subjective scoring of the deposition of silver grains in the autoradiograms.

Renin Immunohistochemistry

Ovarian sections (5 µm) were cut and mounted on SuperFrost/Plus slides (Menzel-Gläser, Braunschweig, Germany), incubated with a polyclonal rabbit antibody raised against mouse renin [29] for 18–20 h at 4°C, and treated by the streptavidin biotin technique (StreptAB-Complex Kit; DAKO A/S, Glostrup, Denmark). The sections were lightly stained with hematoxylin in order to assist localization. The juxtaglomerular cells in cattle kidney and sections treated without the primary antibody served as positive and negative controls, respectively.

Measurement of Protein Concentration

The protein concentration in the cell membrane fractions was determined by the method of Lowry et al. [30].

Statistical Analysis

The data were analyzed by using the Stat-100 statistical analysis package (version 1.29; Biosoft). Nonparametric statistics were used for statistical analysis. Values are given as the median with the range in parentheses. A P value of less than 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Angiotensin II Receptor Analysis in Cell Membrane Fractions

The Scatchard plots were best fitted with a one-site fit (data not shown). The Ang II receptor density was lower in diestrus ovaries compared with proestrus/estrus ovaries those with follicular cysts (Table 1). The dissociation constant, Kd, and the proportion between AT1 and AT2 receptors did not differ in proestrus/estrus, diestrus, or in ovaries with follicular cysts (Table 1).


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  		TABLE 1. Ang II receptor analysis in bovine ovary proestrus/estrus, diestrus, and follicular cysts

Localization of Angiotensin II Receptors

For receptor autoradiography, ovaries of the proestrus (n = 3), estrus (n = 2), postestrus (n = 3) and the diestrus periods (n = 2), as well as two ovaries of pregnancy (Day 75 and Day 140) and two ovaries with follicular cysts were used. Binding of 125I-[Sar1-Ile5-Ile8]-Ang II was found in all ovaries and was highly specific, as revealed by the low, uniform, and nonspecific binding shown by displacement studies using unlabeled [Sar1-Ile5-Ile8]-Ang II (Figs. 1d, 2d, and 3d). Displacement studies with losartan, PD123319, or both showed that both the two Ang II receptor types (AT1 and AT2) were present in the bovine ovaries. No non-AT1/non-AT2 binding sites were found.



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  		FIG. 1. Autoradiograms showing the distribution of 125I-[Sar1-Ile5-Ile8]-Ang II binding to parallel cryosections of an ovary in estrus. Dark regions indicate high density of Ang II receptors. a) AT1 receptor binding. b) AT2 receptor binding. c) Total binding. d) Background labeling. F, Follicle; Fd, dominant follicle. Bar = 1 mm

Follicles In proestrus and estrus, the most intense binding was observed around the dominant follicle (Fig. 1). PD123319 displaced most of the 125I-[Sar1-Ile5-Ile8]-Ang II binding (Fig. 1a), whereas losartan only slightly affected this binding (Fig. 1b), indicating that the Ang II receptor was predominantly type AT2. In the dominant follicle, as in 76% of the other antral follicles, most binding occurred in the theca externa, with a less-intense binding in the theca interna (Fig. 2). Binding in the granulosa layer was not observed (Fig. 2). The other 24% (mainly smaller antral follicles) expressed either few AT1 and AT2 receptors in the same layers or did not express Ang II receptors whatsoever. Around the follicular cysts, intense AT2 receptor binding was located mainly in the theca externa (data not shown). In the primordial, primary, and secondary follicles, no significant labeling was observed.



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  		FIG. 2. Detail of a follicle from parallel cryosections of an ovary in diestrus. a–d) Brightfield and darkfield views of autoradiograms showing the distribution of 125I-[Sar1-Ile5-Ile8]-Ang II binding. Bright grains indicate the presence of receptor. a) AT1 receptor binding. b) AT2 receptor binding. c) Total binding. d) Background labeling. e) Brightfield view of a section stained with hematoxylin and eosin. G, Granulosa layer; TI, theca interna; TE, theca externa; ST, stroma; asterisks, blood vessel. Bar = 75 µm

Corpus luteum In the corpus hemorrhagicum (data not shown) and corpus luteum, the dominance of AT2 receptors was evident in the capsule and connective tissue infoldings (the area of the theca externa; Fig. 3). With progressing age of the corpus luteum, the receptor density declined. In the luteal tissue, the expression of the Ang II receptor was absent. In the corpus luteum of pregnancy, no binding was found, either in the luteal tissue or in the adjacent tissue. No binding was found in the tissue of the corpus albicans (data not shown).



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  		FIG. 3. Autoradiograms showing the distribution of 125I-[Sar1-Ile5-Ile8]-Ang II binding to parallel cryosections of an ovary in diestrus. Dark regions represent high density of Ang II receptors. a) AT1 receptor binding. b) AT2 receptor binding. c) Total binding. d) Background labeling. CL, Corpus luteum; F, follicle; arrow, connective tissue infolding into the CL. Bar = 1.6 mm

Stroma In the stroma, the dominant receptor type was AT1 (Fig. 1) and it showed no changes in relation to the estrus cycle. The density was highest in the stroma adjacent to the tunica albuginea and the stroma surrounding antral follicles (Fig. 1). Strong AT1 receptor binding also occurred in the stroma around the rete ovarii (data not shown). No detectable labeling was observed over the surface epithelium and tunica albuginea. In larger vessels, the surrounding stroma revealed AT1 receptor binding, whereas no apparent radioligand binding was observed in the vessels themselves (Fig. 2, a–c). In the ovaries with follicular cysts, the AT2 receptor dominated in the stroma, and was especially dense between adjacent cystic follicles.

Measurements of ACE Activity

ACE activity was detected in all membrane fractions of ovarian tissue and in follicular fluid, and was completely inhibited by lisinopril or captopril (data not shown). In the follicular fluid from the proestrus/estrus period the ACE activity was 1.3 µmol/min per liter (range 1.0–1.8, n = 5) and in the postestrus/diestrus period it was 1.1 µmol/min per liter (range 0.8–1.3, n = 5). These values are similar to the activity found in bovine serum (unpublished data). In cell membrane fractions of pools of antral follicles, ACE activity was 23.3 µmol/min per gram of membrane protein (range 10.5–78.3, n = 9). In ovaries in proestrus/estrus, the activity was 21.7 µmol/min per gram of membrane protein (range 10.3–41.7, n = 8), and in ovaries with follicular cysts it was 25.0 µmol/min per gram of membrane protein (range 16.7–43.3, n = 7). In tissue of dissected corpus luteum the activity was 2.3 µmol/min per gram of membrane protein (1.3–3.2, n = 7), and in tissue of dissected corpus luteum of pregnancy the value was 9.3 µmol/min per gram of membrane protein (range 2.7–21.7, n = 6).

Localization of ACE

For autoradiographic localization of ACE, ovaries in proestrus (n = 4), estrus (n = 1), postestrus (n = 3), and diestrus (n = 5), as well as ovaries of pregnancy (n = 5; Days 60, 64, 107, 178, and 232) and those with follicular cysts (n = 3) were used. Most ACE binding sites were localized on endothelial cells of stromal vessels (Fig. 4, a and d). Furthermore, some corpora lutea expressed ACE in microvascular endothelial cells localized in the connective tissue infoldings. No binding was observed on granulosa, thecal, or luteal cells (Fig. 4a). Around certain follicles, microvascular endothelial cells displayed binding sites (Fig. 4a). No differences in the binding pattern were observed in ovaries of pregnancy and in ovaries with follicular cysts.



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  		FIG. 4. Autoradiograms showing a) binding of 125I-MK351A to ACE in an ovary of pregnancy (Day 178) and c) background labeling. b) Brightfield view of the respective parallel section stained with hematoxylin and eosin. d) Detail of the stroma of an ovary in diestrus showing binding to microvascular endothelial cells. F, Follicle; CL, corpus luteum, BV, blood vessel. a–c) Bar = 2.1 mm (shown in b). d) Bar = 12.3 µm

Localization of Renin

For immunohistochemistry, ovaries in proestrus (n = 3), estrus (n = 1), postestrus (n = 2), and diestrus (n = 5); ovaries of pregnancy (Days 62 and 107); and one ovary with follicular cysts were used. Renin immunoreactivity was observed in the antral follicles both in granulosa and thecal cells (Fig. 5a), whereas no reactivity was seen in primordial, primary, or early secondary follicles. Also in follicular cysts, no reactivity was observed in these layers. In the corpus luteum, the reactivity was always stronger than in the follicular wall. Both small and large luteal cells showed renin immunoreactivity with varying intensity (Fig. 5c). The same pattern was observed in the corpus graviditas. In the ovarian stroma, the strongest renin immunoreactivity was observed in solitary macrophage-like cells mostly placed near blood vessels (Fig. 5e).



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  		FIG. 5. Immunohistochemical localization of renin in a) an antral follicle, c) a corpus luteum, and e) stroma, and b and d showing negative controls. G, Granulosa layer; TL, thecal layer; ST, stroma; CL, corpus luteum; BV, blood vessel; arrows, small luteal cells; arrowheads, large luteal cells; asterisks, macrophage-like cells. a–d) Bar = 25 µm (shown in b and d). e) Bar = 10 µm


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study confirms the presence of both AT1 and AT2 receptors in the bovine ovary, whereas no non-AT1/non-AT2 receptors could be recognized, as was previously found by our group in the bovine uterus and placenta [21, 23]. Higher Ang II receptor densities were found in cell membrane fractions of ovaries in estrus than in diestrus, providing strong evidence for a cyclic variation of the expression of these receptors. In concert with this finding, autoradiography revealed the strongest marking in the dominant follicle of the proestrus and estrus periods.

Most Ang II receptors in antral follicles were localized in the theca externa, whereas their expression was sparse in the theca interna. No expression was observed in the granulosa cell layer. This is in accordance with a study on isolated bovine granulosa, thecal, and luteal cells, in which Ang II binding sites were found only on thecal cells [9]. The individual follicles of an ovary will be at different maturation stages, some belonging to the pool of growing follicles and some belonging to the group of atretic follicles [31, 32]. This could explain the difference in Ang II receptor expression observed by autoradiography, in which 24% of smaller antral follicles expressed no or sparse with AT1 as the predominating type, while the larger follicle (also in the ovary of pregnancy) always expressed many receptors, predominantly those of AT2. In the bovine ovary, the density of Ang II receptors was positively correlated with follicular diameter and tissue weight, but did not correlate with follicular fluid steroids [8]. In a recent study, the level of AT2 receptor mRNA dropped transiently in bovine follicles with estradiol concentrations ranging from 20–180 ng/ml (mean follicle diameter = 12.6 mm), but increased again with estradiol concentrations greater than 180 ng/ml (mean follicle diameter = 14.4 mm) [20]. This study is difficult to compare with our results because the mRNA was measured in dissected theca interna only. As the present study clearly demonstrates, Ang II receptors are expressed primarily in the theca externa.

The fact that Ang II receptors were apparently absent in luteal cells is in accordance with a previous study in cell membrane fractions of bovine luteal cells [9]. The Ang II receptors observed in the connective tissue infoldings of the corpus luteum explain the expression of Ang II receptors previously demonstrated using total bovine corpus luteum tissue [8]. Connective tissue cells could also be responsible for the increased expression of the bFGF mRNA observed after stimulation of a primary cell culture of bovine corpus luteum cells with Ang II [18].

Ang II is known to regulate blood-flow and vascular permeability [33], but as in the bovine uterus and placenta [21, 23], little or no Ang II receptor expression was observed in the endothelial and muscle cells of the ovarian blood vessels. However, Ang II may nevertheless still contribute to this local vascular regulation.

The highest density of both Ang II receptor types was observed around the antral follicles and the corpus luteum. Ang II has angiogenic properties and growth-promoting effects through its AT1 receptor [34, 35], while it can mediate antiangiogenic properties and apoptosis through the AT2 receptor [35, 36]. The colocalization of both receptor types in these tissues suggests that Ang II may participate in the regulation of angiogenesis and growth through cross-talk between the receptor types as suggested by Inagami et al. [37]. In this way, Ang II may contribute to follicular and luteal development. A further definition of the morphological and biochemical characteristics of the individual follicles and their Ang II binding pattern could improve our understanding of the putative role of Ang II in folliculogenesis.

Both renin and ACE enzymes, which are responsible for the conversion of ANG to Ang II were demonstrated in the bovine ovary, indicating that Ang II may be produced locally. ACE was demonstrated both in aspirated follicular fluid and in ovarian tissue. By autoradiography, we localized ACE on microvascular endothelial cells and endothelial cells of larger blood vessels. This is in concert with a recent study that suggested the presence of ACE on microvascular endothelial cells isolated from developing bovine corpus luteum [19]. The presence was indicated by the conversion of Ang I to Ang II by these cells, a reaction that was inhibited by the ACE inhibitor captopril. Also, in the rat ovary, ACE was demonstrated in blood vessels [38, 39], and furthermore, on the germinal epithelium, an outer rim of certain corpus luteum, and on granulosa cells of certain follicles [38, 39]. In the cow, a correlation between ACE activity and the progesterone concentration in the follicular fluid was found (personal communication with A.H. Nielsen). Because we found no ACE in the granulosa layer and high progesterone concentrations mostly reflects follicular atresia, it could be speculated that ACE activity detected in the follicular fluid originates from endothelial cells in the thecal layers and thereby reflects the condition of the basal lamina.

Renin immunoreactivity was found in both granulosa and thecal cells of all antral follicles. In human and rat ovaries, thecal cells stain strongly in all follicles, whereas granulosa cells stain heavily only in preovulatory follicles [40]. As recently as 1989, bovine follicles were shown to possess renin-like activities with thecal cells as the major source [41]. An earlier study by our group suggested that the wall of atretic follicles contained less renin, because the renin concentrations in the follicular wall correlated negatively with progesterone concentrations in the follicular fluid [42]. However, the immunohistochemical staining could not reflect this variation.

It is well known from rats, cows, pigs, and humans that luteal tissue contains renin [4, 10, 43]. In the present study, all luteal cells showed renin immunoreactivity and the corpus luteum always stained more strongly than the follicles. This is in contrast to an earlier finding showing that the active renin concentration in the bovine corpus luteum was similar to that in the follicle walls [10]. The discrepancy may reflect the difficulty of comparing results obtained by different methods. When measuring active renin, the concentration in homogenated tissue—composed of many different cell types—is measured. Immunohistochemistry allows localization of renin to single cell types, but visual quantitation is more uncertain.

In the bovine ovarian stroma, solitary cells staining strongly for renin and placed adjacent to blood vessels were identified. Immunohistochemical studies combined with high-performance liquid chromatography or double immunostaining have shown both human and rat macrophages and monocytes to be renin-containing cells [44, 45]. Due to the localization and shape of the bovine solitary cells and the fact that the bovine ovary contains many macrophages, they are suggested to be macrophages.

The fact that Ang II receptor expression was high in ovaries with follicular cysts and that no renin was observed in the wall of the cystic follicles could indicate that disturbances in the RAS may lead to follicular cyst formation.

Although the RAS in the bovine ovary could affect oocyte maturation, ovulation, and expression of vascular modulators and gonadal steroids, its role and significance are still unclear. The relative merits of the present study still need to be assessed. Elucidation of the bovine ovarian RAS may contribute to a better understanding of the development and selection of follicles and thereby contribute to the development of an improved system for in vitro oocyte maturation technique. Furthermore, our results stress the importance of studying this local system species specificity, and this necessarily should lead to investigations on human ovary and the influence that treatment with ACE inhibitors may have.

Our findings emphasize that although there are similarities, there is a considerable species variation in the distribution of the components of the ovarian RAS, which could reflect species differences in the regulation of folliculogenesis. In the bovine ovary, both receptor types were present in the periphery of the follicles and the corpus luteum. Therefore, it is suggested that Ang II may contribute to follicular and luteal development. In conclusion, our findings support the concept of an active and regulated renin-angiotensin system in the bovine ovary.


   ACKNOWLEDGMENTS
 
We are grateful to Prof. Poul Hyttel, D.V.M., Ph.D., D.V.S.C.; Prof. Niels Henrik Diemer; and laboratory technician Marianne Nielsen for their assistance and helpful suggestions. The skillful technical assistance of Christina T. Larsen, Susanne Holm, Iben Thomsen, and Hanne Holm is gratefully acknowledged.


   FOOTNOTES
 
First decision: 1 May 2001.

1 This study was partially supported by grants from the Danish Agricultural and Veterinary Research Council, the Nordic Insulin Fund, and the NOVO Fund. Losartan (DuP 753) and PD123319 were gifts from DuPont Merck Pharmaceutical Company, Wilmington, DE, and Parke-Davis, Ann Arbor, MI, respectively. Back

2 Correspondence: K.H. Schauser, Department of Anatomy and Physiology, The Royal Veterinary and Agricultural University, Grønnegaardsvej 7, DK-1870 Frederiksberg C, Denmark. FAX: 45 35282525; ks{at}kvl.dk Back

Accepted: July 16, 2001.

Received: March 30, 2001.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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