Regulation of tight junction proteins occludin and claudin 5 in the primate ovary during the ovulatory cycle and after inhibition of vascular endothelial growth factor
1Department of Obstetrics and Gynecology, University of Ulm, Prittwitzstrasse 43, 89075 Ulm, Germany
2Medical Research Council Human Reproductive Sciences Unit, The Quee
s Medical Research Institute, Edinburgh EH16 4TJ, UK
3 Correspondence address. E-mail: webmaster{at}tine-wulff.de
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Abstract |
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Ovarian follicular and corpus luteum development, including angiogenesis, are characterized by cell–cell rearrangements that may require dynamic changes in cell–cell adhesion. The present study investigates the expression of tight junction proteins occludin and claudin 5 during follicular and luteal development in the primate ovary and after inhibition of vascular endothelial growth factor (VEGF) by VEGF trap treatment. Occludin was localized to the plasma membrane of granulosa cells. During follicular development occludin staining decreased significantly (P < 0.05) and disappeared completely by the ovulatory stage. After inhibition of VEGF, occludin staining was significantly (P < 0.05) higher in the granulosa of secondary and tertiary follicles compared with controls. Claudin 5 was exclusively localized to the theca vasculature. A significant (P < 0.05) increase in staining was detected from the pre-antral to the antral and ovulatory stage. However, dual staining with CD31 revealed that within the theca endothelium the amount of claudin 5 remained constant during follicular development. Treatment with VEGF trap throughout the follicular phase revealed a lack of claudin 5 staining in the theca interna but no difference was observed in the remaining theca externa vasculature. In the corpus luteum, claudin 5 was also localized in the vasculature. Treatment with VEGF trap in the mid-luteal phase resulted in a significant increase in staining (P < 0.05). These results led us to hypothesize that tight junctions are involved in regulation of follicular growth, antrum transition and follicular angiogenesis which is compromised by VEGF inhibition. VEGF may influence luteal vascular permeability by regulation of the endothelial specific tight junction protein claudin 5.
Key words: ovary/permeability/primate/tight junctions/VEGF trap
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Introduction |
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In the ovary, dynamic morphological changes occur during follicular growth and corpus luteum development. Processes such as granulosa proliferation, antrum formation and theca development during follicular growth, as well as angiogenesis during follicular and corpus luteum development, are dependent on dynamic cellular rearrangements. Cell–cell adhesion plays a role in regulating these processes as an essential component for communication between the adjoining cells (Bazzoni and Dejana, 2004; Walz et al., 2005) to ensure a specific tissue morphology and function. The phenomenon that an epithelial or endothelial cell stops proliferation as soon as it comes in contact with another cell is known as contact inhibition. This process has an important role in controlling cellular growth and is mediated by adhesion proteins (Lampugnani et al., 2006). Thus, a tissue or a tissue compartment that continues to undergo remodelling, such as the ovary, may require tightly controlled dynamic changes in localization and expression of adhesion molecules.
Major components of intercellular junctions are tight junctions (Dejana, 2004; Schneeberger and Lynch, 2004) which are composed of different transmembrane proteins promoting homophilic interaction. The cytoplasmatic domain of the transmembrane adhesion molecules bind to linker proteins that in turn anchor the adhesion complex to the cytoskeleton. Of these molecules, occludin and the claudins are the most extensively studied. Although occludin is a highly conserved molecule, claudins comprise a family of over 20 different proteins, some of which, such as claudin 5 in endothelial cells, are expressed in a tissue specific manner (Morita et al., 1999, 2003; Peppi and Ghabriel, 2004). Tight junctions are the most apical component of the intercellular junctional complex which completely seal the space between neighbouring cells. Thus, they are not only important for cell communication but also serve for providing a ‘barrier’ and a ‘fence’ within the membrane by regulating paracellular permeability and maintaining tissue and cellular integrity.
Recently, we began to study the differential adhesion protein expression during remodelling in the human corpus luteum (Groten et al., 2006). Here, we investigate localization and changes of tight junction proteins during normal follicular development in the marmoset ovary. We have also demonstrated in the marmoset model that inhibition of vascular endothelial growth factor (VEGF) has a major inhibitory effect on ovarian angiogenesis, development and function (Wulff et al., 2001b, 2002; Taylor et al., 2007). In the current study, we address the question of whether some of these effects may be mediated via alterations in the interaction between VEGF and adhesion proteins. Cell adhesion proteins have been shown to interact with signalling pathways of growth factor receptors and vice versa. VEGF is directly involved in tyrosine phosphorylation of junctional proteins (Esser et al., 1998) resulting in down-regulation of tight junction proteins (Ghassemifar et al., 2006). A direct effect of VEGF was shown in HepG2 cells in which VEGF induces disruption of occludin-delineated tight junctions (Schmitt et al., 2004). We therefore investigated whether VEGF-inhibition interferes with expression of follicular tight junctions that may contribute to the inhibition of follicular development in vivo. Furthermore, it has been demonstrated that VEGF supressed tight junctions in cultured endothelial cells (Ghassemifar et al., 2006), decreased transendothelial resistance and caused changes in actin fibers in HUVECs indicating a decreased endothelial permeability (Villasante et al., 2007).
Recently, we have shown that inhibition of VEGF in the ‘post-angiogenic’ period in the corpus luteum i.e. during mid- to late-luteal phase resulted in a rapid, marked and sustained suppression of plasma progesterone levels which is unexplained (Fraser et al., 2006). We therefore tested the hypothesis that VEGF inhibition may induce rapid changes in expression of endothelial claudin 5 which may compromise luteal vascular permeability.
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Material and Methods |
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VEGF trap
The VEGF trap is a recombinant chimeric protein comprising portions of the extracellular, ligand binding domains of the human VEGF receptors Flt-1 (VEGF-R1, Ig domain 2) and KDR (VEGF-R2, Ig domain 3) expressed in sequence with the Fc portion of human IgG as described in detail previously (Wulff et al., 2002).
Animals
Ovaries that were studied had been obtained from previous investigations (Wulff et al., 2002; Fraser et al., 2006; Taylor et al., 2007) in which VEGF trap was administered to marmosets between Days 0 and 10 of the follicular phase or at the mid-luteal phase for 1 day. Adult female common marmoset monkeys (Callithrix jacchus) with a body weight of approximately 350 g and regular ovulatory cycles (28-day cycle length) were housed as described previously (Fraser et al., 1999). Blood samples were collected three times per week by femoral venipuncture, and plasma was subjected to progesterone assay as described previously to determine the time of ovulation (Wulff et al., 2001c).
Treatment
Experiments were carried out in accordance with the Animals (Scientific Procedures) Act, 1986, and were approved by the local ethical review committee. To synchronize timing of ovulation, marmosets were given prostaglandin F (PGF2)
analogue (Cloprostenol, Planate, Coopers Animal Health Ltd, Crewe, UK), intramuscularly on Days 13–15 of the 20-day luteal phase to induce luteolysis = cycle Day 0. VEGF trap was administered beginning at the time during follicular recruitment to inhibit follicular angiogenesis and development. Eight marmosets were treated with VEGF trap at a dose of 25 mg/kg, injected s.c. on Days 0, 2, 4, 6 and 8 of follicular phase (n = 4) (Wulff et al., 2002) or as a single injection of 25 mg/kg on Day 0 (n = 4) (Taylor et al., 2007). Ovaries were collected on Day 10 of the cycle. As responses were similar, results from both groups were combined. Ovaries from four controls containing pre-ovulatory follicles were collected on Day 10 after prostaglandin treatment. Control animals were treated with vehicle, containing 5 mM phosphate, 5 mM citrate, 100 mM sodium chloride, 0.1% (wt/vol) Tween20 and 20% (wt/vol) sucrose.
The second experiment was designed to determine the effects of VEGF inhibition on the corpus luteum in the ‘post-angiogenic’ period of luteal phase that might compromise luteal permeability (Fraser et al., 2006). Marmosets were treated with a single injection of 25 mg/kg s.c. VEGF trap at the mid-luteal phase. Ovaries were collected 1 day later. Control animals received 25 mg/kg s.c. Fc portion of human IgG at the mid-luteal phase (n = 4 per group).
Ovaries were collected and fixed in 4% neutral buffered formalin as described (Fraser et al., 2006). After 24 h, the ovaries were put into 70% ethanol, dehydrated and embedded in paraffin according to standard procedures.
Tissue processing and haematoxylin and eosin staining
The embedded ovaries were sectioned (5 µm) and tissue sections were placed onto BDH SuperFrost slides (BDH, Merck & Co., Inc., Poole, UK). Consecutive sections were used for H&E, occludin, claudin 5 and CD31 staining. Tissue sections were dewaxed in xylene, rehydrated in descending concentrations of ethanol and washed in distilled water. For morphological classification of the follicles sections were stained with haematoxylin (Richard-Allan, Richland, MI, USA) for 5 min, followed by a wash in water and acetic alcohol before staining with eosin (Richard-Allan) for 20 s. After dehydrating in ascending concentrations of ethanol and xylene, sections were mounted.
Immunhistochemistry
To localize the tight junction proteins occludin and claudin 5, the following antibodies were used: rabbit anti-human occludin (Zymed, San Fransisco, USA, 71-1500; 1:100 dilution) and mouse anti-human claudin 5 (Zymed 18-7364; 1:100 dilution). CD31 staining was performed to identify the endothelial compartment using a mouse anti-human CD31 antibody (Dako Corp., Copenhagen, Denmark, 1:100 dilution). Negative controls were performed for all antibodies by replacing the first antibody with Tris-buffered saline (TBS). Antigen retrieval was achieved by pressure cooking (Clypso pressure cooker, Tefal, Essex, UK) in 0.01 mol/l citrate buffer, pH 6 for 7 min. Sections were left for 20 min in hot buffer before cooling in TBS.
For occludin and claudin 5 staining, the first antibody was applied for 15 min at room temperature in 2% normal rabbit serum (NRS). Signal amplification was carried out using the CSA-System Peroxidase (Dako, K1500). The instructions of the manufacturer were followed precisely.
CD31 and claudin 5 dual staining
Immunofluorescence double-staining was performed using the TSA-Kit (Perkin Elmer, Boston, USA). Sections of paraffin-embedded tissue were dewaxed and rehydrated using xylol and ethanol, respectively, and transferred to TN-buffer. Slides were cooked under pressure in 10 mmol/l of citrate buffer (pH 6.0) for 7 min, left in the hot buffer for another 20 min and transferred to TN-buffer for 5 min. Endogenous peroxidase was quenched for 30 min in 180 ml methanol + 20 ml H2O2 (30%), and slides were again transferred to TN-buffer for 5 min. After pre-incubation with avidin and biotin (Serotec, Oxford, UK), 8 drops per ml blocking serum (2.5 g bovine serum albumin per 40 ml TN-buffer + 20% NRS), for 30 min, respectively, the slides were incubated with a dilution of 1:100 mouse anti-Claudin 5 antibody at 4°C over night. The slides were washed in TN-buffer with 0.1% Tween for 3 x 5 min, followed by incubation with biotinylated secondary antibody (rabbit anti-mouse, 1:300, Dako) for 45 minutes. Washing in TN-buffer + Tween and incubation with SA-horse-radish peroxidase (HRP) and Fluoresceine tyramide was performed according to the instructions of the manufacturer. Incubation with the second primary antibody mouse anti-CD31 (1:30, Dako) over night at 4°C was followed by washing, incubation with secondary antibody, SA-HRP and TMR tyramide as described above. Mounting was performed with Mowiol.
Analysis of data
Quantitative analyses were performed using an Olympus BH2 microscope, Spot Insight QE camera and Image-Pro Plus version 4.5 for Windows (Media Cybernetics, Silver Spring, MI, USA).
Morphological characterization of ovarian follicles
Consecutive sections stained for haematoxylin and eosin were used to classify follicular stages. Stages of follicular development were defined as follows: primary follicles (containing only one granulosa cell layer), early secondary follicles (two to four granulosa cell layers), late secondary follicles (more than four granulosa cell layers, no antrum), tertiary follicles (follicles containing an antrum) and ovulatory follicles (large antral follicles >2 mm diameter). Follicles were classified as healthy if they contained a normal-shaped oocyte surrounded by granulosa cells that were regularly apposed on an intact basement membrane with normal appearance of granulosa cell nuclei without signs of pycnosis. Follicles not fulfilling these criteria were classified as unsuitable for analyses. Only follicles with a visible oocyte containing a nucleus were considered to ensure proper follicular classification.
Quantification of immunocytochemistry
Quantification was carried out by two different observers blinded for the staining and the treatment. Four representative cross sections of the ovaries per animal were subjected to morphometric analyses each for occludin, claudin 5 and CD31 staining, respectively.
In total, 106 follicles were analysed: 44 secondary, 49 tertiary and 13 ovulatory follicles. Area of positive staining in the follicles for occludin was measured at x200 magnification. In all follicles, the whole cross-sections were analysed. Captured images were thresholded, and the granulosa cell compartments outlined and analysed. The positive staining for occludin was then calculated per unit area of the granulosa compartment and expressed as a mean value for the number of follicles assessed within each follicular stage and per animal.
To analyse the percentage of claudin 5 in the theca endothelium, a dual staining for CD31 and claudin 5 was performed. To compare different antibody staining, grid counting proved to be optimal. The theca compartment was outlined and a grid superimposed. The size of the single grid box was 5 µm2. The number of positive hits (i.e. when the staining hits a crossing point on the grid) was counted, as well as the total number of crossing points on the grid within the outlined theca compartment. The percentage of positive hits in the theca was calculated for each antibody by dividing the number of positive hits by the total number of crossing points. Values were expressed as means for the number of follicles within each follicular stage and per animal. Furthermore, the percentage of positive claudin 5 staining within the theca endothelium was calculated by division of percentage of claudin 5 hits by percentage of CD31 hits and expressed as a mean value for the number of follicles assessed within each follicular stage and per animal.
Quantitative image analyses for claudin 5 staining in the corpus luteum were performed in the whole cross sectional area of the corpus luteum. All corpora lutea were analysed under x40 magnification for all parameters. A grid of 1302 crossing points was superimposed, and the number of positive hits in the whole captured image for claudin 5 and CD31 staining was counted and the percentage of claudin 5 within the luteal vasculature estimated. Single values for individual animals were calculated by taking the mean of the measurements obtained on all the corpora lutea in that animal.
Statistical analysis
Data obtained for different follicular stages were tested for significant differences using ANOVA, followed by Duncan's multiple range test. Effects of different treatments compared with controls were determined using a two-tailed, unpaired t-test. Differences were considered to be significant at P < 0.05. The tests were performed using SPSS version 6.1 for Macintosh (SPSS, Inc., Chicago, IL, USA). All values are given as the mean ± SEM.
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Localization of occludin in the normal cycle
In the ovary, occludin was present in follicles and in the endothelium of large stroma vessels (Fig. 1). It was not present in the corpus luteum. Within the follicles, occludin was localized to the granulosa cells and the plasma membrane of the oocyte (Fig. 1a, inset). It was absent in theca cells and the theca endothelium. In early follicular stages, i.e. primary and early secondary follicles, occludin staining was present in the plasma membrane of the granulosa cells (Fig.1a, inset). A gradual decrease in occludin staining within the granulosa was notable during follicular growth. In early to late secondary follicles, occludin staining disappeared from the granulosa layers in the vicinity of the oocyte (Fig. 1b). With increasing follicular size, staining became less strictly localized to the plasma membrane whereas cytoplasmic staining was more evident. By the antral stage, occludin staining remained localized in the plasma membrane only in the granulosa cells lining the basement membrane (Fig. 1c). In pre-ovulatory follicles, occludin staining was completely absent in granulosa cells (Fig. 1d). Image analyses revealed a significant continuous decrease (P < 0.05) of occludin (Fig. 1e) staining during follicular development from primary (41.3 ± 2.3%) to early secondary (33.1 ± 0.6%), late secondary (29.7 ± 1.4), tertiary (25.0 ± 0.4%) and ovulatory stage (0%).
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Changes of occludin after VEGF inhibition
After VEGF inhibition, occludin staining was still present in the vicinity of the plasma membrane of granulosa cells in early follicular stages (Fig. 2a and b). However, in later stages, occludin staining appeared to be localized in the cytoplasm after VEGF inhibition (Fig. 2c and d). The gradual decrease of occludin staining was less pronounced after treatment. Measurements revealed only a significant decrease of occludin staining (P < 0.05) if primary follicles were compared with tertiary follicles (Fig. 2e). Image analyses further revealed that levels of occludin staining for all follicular classes was always higher after treatment than compared with the control (Fig. 2e) which was significant in late secondary and tertiary follicles.
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Localization of claudin 5 in the normal cycle
Claudin 5 is an endothelial specific tight junction protein, and we found it exclusively localized to the ovarian vasculature. In follicles, claudin 5 was detectable in the theca vasculature as soon as the follicle has established a capillary system, i.e. by the late secondary stage (Fig. 3a). With increasing follicular size, the staining for claudin 5 was more evident (Fig. 3b), especially in pre-ovulatory follicles (Fig. 3c). No difference was found between healthy (Fig. 3b) and atretic follicles (Fig. 3d). During the early luteal phase, claudin 5 was present in the theca capillaries that began to penetrate into the granulosa (Fig. 3e). By the mid-luteal phase, claudin 5 staining was evenly distributed throughout the luteal vasculature (Fig. 3f). Image analysis (Fig. 4g) revealed a significant increase in claudin 5 staining in the theca from late secondary (4.14% ± 0.1%) to tertiary (7.96% ± 0.2%) and to the ovulatory stage (15.3% ± 0.4) (data not shown in Fig. 4g). The same pattern was found for CD31 staining. Dual staining for CD31 and claudin 5 revealed a constant amount, 80% ± 6%, of claudin 5 staining within the endothelium of all follicular classes.
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Changes of claudin 5 in follicles after VEGF inhibition
In contrast to controls (Fig. 4a, c and e), claudin 5 staining was completely absent in the theca interna of late secondary and tertiary follicles (Fig. 4b, d and f) after VEGF trap treatment. Staining for claudin 5 was only found in theca externa vessels. Image analyses confirmed a significant decrease of claudin 5 staining in the theca in late secondary and tertiary follicles after VEGF trap (Fig. 4e). However, when comparing the percentage of positive endothelial staining for claudin 5, no significant difference was detected in late secondary and tertiary follicles after treatment. The total amount of claudin 5 in the theca externa vessels was constant (i.e. 80% ± 8.1%).
Changes of claudin 5 in the corpus luteum of the normal cycle and after VEGF inhibition
Claudin 5 was localized to the luteal endothelium and evenly distributed during mid-luteal phase when the corpus luteum and its vasculature network reached functional maturity (Fig. 5a, c and e). After VEGF inhibition (Fig. 5b, d and f) claudin 5 remained evenly distributed within the luteal vasculature, but the amount of claudin 5 staining in the endothelium appeared to increase (compare Fig. 5e with f). Quantitative analyses confirmed a significant increase of claudin 5 staining in the endothelium after VEGF inhibition (Fig. 5g).
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Discussion |
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In this study, we demonstrate for the first time in a primate species that the tight junction proteins occludin and claudin 5 are differentially expressed during follicular development in the ovary. Furthermore, we show that suppression of follicular angiogenesis by VEGF inhibition significantly affected occludin expression in the follicular granulosa cells, indicating disturbance of cell–cell communication may contribute to suppression of follicular development (Wulff et al., 2002). These observations led us to hypothesize that cell–cell adhesion mediated by tight junction proteins occludin and claudin 5 may play a critical role in regulating physiological tissue remodelling processes in the ovary.
During follicular development granulosa cells proliferate, contributing to the increase in size of the follicle (Jablonka-Shariff et al., 1996; Hirshfield, 1997). The granulosa cell–cell rearrangement that continues throughout follicular maturation may involve changes in cell adhesion. Here, we observed a decrease of occludin staining in the plasma membrane of granulosa cells from the late secondary to pre-ovulatory follicles, at which stage staining was completely absent. Tight junctions are involved in cell sealing which will influence the regulation of extracellular fluid transport between cells. The loss of occludin during follicular development may be involved in antrum formation, since antrum formation must involve alteration of granulosa cell–cell adhesion and sealing. In accordance with our observations, immunoblot studies in rat and mouse granulosa cells of antral follicles showed a decrease of adhesion junctional proteins (Sundfeldt et al., 2000; Kawagishi et al., 2005). After VEGF inhibition, occludin was maintained and image analyses revealed higher staining after treatment when compared with controls. However, morphologically, the former membrane-associated staining became less obvious, whereas cytoplasmic staining was evident. Thus, the increase shown by image analyses is likely to result from an increase in cytoplasmic staining. However, occludin in the cytoplasm is non-functional (Alexander and Elrod, 2002), indicating that inhibition of normal vascular development severely affected granulosa cell–cell adhesion and communication by disruption of occludin delineated tight junctions. In previous studies (Wulff et al., 2002; Taylor et al., 2007) we showed that VEGF inhibition from Days 0 to 10 of the follicular phase resulted in suppression of follicular development, granulosa proliferation, prevention of antrum formation and ovulation which further underlines the role of occludin in regulation of antrum transition. Since marmoset granulosa cells do not appear to exhibit VEGF receptors, the disturbance of follicular growth is secondary to the treatment. Having prevented the development of a functional vasculature, the granulosa is deprived of nutrient supply so that the granulosa cells degenerate and switch off all cellular signalling. This may also affect tight junction proteins and cell–cell communication, thus the normal remodelling processes during follicular development.
Occludin staining was further evident in the plasma membrane of the oocyte of all follicle classes. The oocyte is surrounded by the zona pellucida but interacts with the granulosa through thin extensions of the granulosa cells penetrating the zona pellucida which connect to the oocyte surface. Thus, occludin may be involved in granulosa–oocyte adhesion and communication. In contrast to the granulosa, occludin staining in the oocyte remained after antrum formation, indicating that occludin further serves for the exchange of metabolic and developmental signals.
Occludin is a tight junction protein that is involved in sealing the intercellular space to serve as a blood tissue barrier. Accordingly, occludin is highly expressed in tight vessels such as brain capillaries (Hamm et al., 2004; Harhaj and Antonetti, 2004). With respect to its localization in the vasculature of the ovary, occludin was only found in the large stroma vessels but not in the follicular or luteal vasculature. Interestingly, VEGF inhibition suppressed follicular and luteal angiogenesis, whereas the stroma vasculature of the ovary was unaffected. Thus, the stroma vessels appear to be more resistant to the treatment and occludin may serve as a protective mechanism.
In contrast, Claudin 5 was detected in the theca vasculature as soon as a capillary system became established by the late secondary stage. Claudin 5 staining significantly increased in the theca from late secondary to the ovulatory follicular stage. Thus, theca vascular development during folliculogenesis requires endothelial cell adhesion necessary for the establishment of a functional vasculature that is mediated by claudin 5 but not occludin. The highest amount of claudin 5 in the theca vasculature was detected in the fully matured ovulatory follicle when the angiogenic process is complete. After ovulation, these theca capillaries are the source of new capillary sprouting which occurs during luteal angiogenesis (Dickson et al., 2001; Wulff et al., 2001c; Fraser and Wulff, 2003; Fraser et al., 2006). However, these early luteal capillaries do not express claudin 5 indicating that during early luteal angiogenesis, suppression of cell adhesion may allow endothelial cells to proliferative and expand. By the mid-luteal phase, when endothelial cell proliferation has declined, claudin 5 is evenly distributed within the luteal vasculature. Here, claudin 5 may have a role in contact inhibition of endothelial cells, thus endothelial cell proliferation decreases and the vasculature becomes stabilized. The role of adhesion proteins for contact inhibition has been demonstrated in cell culture studies (St Croix et al., 1998; Grazia Lampugnani et al., 2003; Bazzoni and Dejana, 2004; Nelson et al., 2004; Noseda et al., 2004; Lampugnani et al., 2006). As soon as endothelial cells come in contact with another cell and adhesion molecules link together, the endothelium becomes less affected by the proliferative stimulus of VEGF. On the one hand, contact inhibition may be a prerequisite for the endothelial cells to form tubules which have to become stabilized by cell adhesion to form new capillaries. On the other hand, in certain areas of a tissue in which endothelial cell proliferation is required to establish a vasculature, the proliferating cells should not adhere to each other. Accordingly, we did not detect any claudin 5 staining in luteal capillaries during early luteal phase, whereas it was found during mid-luteal phase in the established vasculature.
That cell adhesion plays an important role in regulation of angiogenesis has been shown by Nakhuda et al. (2005) who demonstrated that inhibition of VE-cadherin is followed by suppression of angiogenesis and degeneration of the vasculature. VEGF is a key molecule in regulation of ovarian angiogenesis (Wulff et al., 2000, 2001a–c, 2002). Furthermore, a close relationship between VEGF and cell adhesion signalling pathways has been demonstrated (Esser et al., 1998). Thus it was rational to assume that in vivo inhibition of VEGF may have affected the protein localization and expression in the ovary. VEGF inhibition in theca vasculature appeared to have down-regulated claudin 5 staining; however, the pattern of claudin 5 staining resembled the staining of CD31, which was in accordance with the observations of our previous studies (Wulff et al., 2002; Taylor et al., 2007). Thus, since the theca vasculature itself was suppressed, claudin 5 staining per theca area had to decrease. Regarding the amount of claudin 5 staining within the remaining theca externa vessels, no quantitative difference was found after VEGF inhibition throughout the whole follicular phase. In contrast, a 1-day treatment with VEGF trap during the mid-luteal phase was associated with a significant increase in claudin 5 within the luteal endothelium. In the former study, we also demonstrated that this treatment was followed by a marked and sustained suppression of progesterone (Fraser et al., 2006). It was suggested that a short-term inhibition during the ‘post-angiogenic’ period of the mid-luteal phase may affect luteal permeability. The current results led us to hypothesize that after VEGF inhibition, the observed claudin 5 increase may result in sealing of the intercellular space which may affect paracellular transport, so that hormone precursors may not reach the luteal cells and/or progesterone may not be released into the blood stream. Thus VEGF may influence luteal permeability by regulation of tight junction proteins.
In conclusion, here we have demonstrated that tight junction proteins are differentially expressed during follicular development which may indicate their key role in regulation of follicular growth, antrum transition and ovarian angiogenesis. Furthermore, this is the first report in a primate species that in vivo inhibition of VEGF leads to disruption of occludin delineated tight junctions in follicles associated with suppression of follicular growth. In addition, it is demonstrated for the first time that in vivo inhibition of VEGF up-regulates claudin 5 protein expression in the corpus luteum which may indicate that VEGF may be responsible for regulation of luteal permeability.
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This work was supported by the Deutsche Forschungsgemeinschaft (DFG).
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We thank Dr J.S. Rudge, Dr S.J. Weigand and Regeneron Pharmaceuticals Inc. (Tarrytown, USA) for expert advice and gift of the VEGF trap, Keith Morris and staff for animal care and Helen Wilson and Regina Konrad for help with histology.
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These authors contributed equally to the manuscript. Disclosure summary: the authors of this manuscript have nothing to disclose. 
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Submitted on June 20, 2007; resubmitted on August 31, 2007; accepted on September 5, 2007.
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