Molecular Human Reproduction, Vol. 7, No. 1, 27-34,
January 2001
© 2001 European Society of Human Reproduction and Embryology
Ovary and oogenesis |
Differential expression patterns of cathepsins B, H, K, L and S in the mouse ovary
1 Departments of Molecular Biology and Medical Biochemistry, and 2 Obstetrics and Gynaecology, Turku University Central Hospital, University of Turku, FIN-20520 Turku, Finland
Abstract
Cathepsins B, H, K, L and S belong to a family of lysosomal cysteine proteinases which participate in a variety of proteolytic processes, including degradation of extracellular matrix. Although the presence of cathepsin mRNAs in the ovary has been reported earlier, very little information is available on their temporospatial expression. In the present study, Northern analysis revealed cyclic changes in the mRNA levels for cathepsins B, H, K, L and S during the 4-day oestrous cycle in the mouse ovary. Immunohistochemical localization revealed distinct expression patterns suggesting different functions for the cathepsins studied. Cathepsin B was predominantly seen in the germinal epithelium throughout the oestrous cycle. Upon follicular maturation, an increasing number of granulosa cells became positive for all cathepsins. Strong cathepsin H staining was sharply defined in theca externa which also stained for cathepsins K and S. Corpus luteum was the predominant location of cathepsin L. The distribution of cathepsin S resembled that of cathepsin L. The developing oocyte stained positive for all cathepsins. In-situ hybridization confirmed the differential production of cathepsin mRNAs by granulosa, thecal and luteal cells. These complex temporal and spatial expression patterns at different stages of the oestrous cycle and follicular development suggest divergent functions for specific cathepsins in follicular development, growth and rupture.
cathepsin/extracellular matrix/mRNA/ovary
Introduction
Cathepsins B, H, K, L and S belong to the gene/protein family of lysosomal cysteine proteinases, whose catalytic activity is based on a cysteine residue in the active site (Rawlings and Barrett, 1994
; Kirschke et al., 1998). All these cathepsins are synthesized as (pre)proenzymes, which are processed into catalytically active proteolytic enzymes of 23–30 kDa. The activity of cysteine proteinases is dependent on pH values of <7, as found in lysosomes, where these enzymes perform their main biological function. However, there is increasing evidence for extracellular functions of cathepsins produced by macrophages, osteoclasts, fibroblasts, and transformed cells into specific pericellular locations where low pH values are observed (Chapman et al., 1997; Kirschke et al., 1998). The resorption lacuna of an osteoclast is an example of such microenvironment where cathepsin K plays an important role in the degradation of bone matrix (Väänänen, 1993; Saftig et al., 1998). The activity of cathepsins is also controlled by their inhibitors, cystatins. An alteration in the cathepsin/cystatin balance may result in uncontrolled proteolysis as seen in inflammatory disorders and during tumour growth (Chapman et al., 1997; Kirschke et al., 1998).
Several studies performed on the tissue distribution of mRNAs coding for specific cathepsins have shown that cathepsins B, H, K, L and S are produced in the human and mouse ovary, often at relatively high levels (Petanceska and Devi, 1992; Brömme and Okamoto, 1995; Rantakokko et al., 1996; Kirschke et al., 1998; Söderström et al., 1999). These studies have not, however, taken into account the physiological functional status of the ovary analysed. Consequently, the role of cathepsins in the ovary remains obscure. Studies on other tissues have shown that cathepsins, together with matrix metalloproteinases (MMPs), play an active role in the degradation of extracellular matrix (ECM), including collagens (Maciewicz and Wotton, 1991; Kakegawa et al., 1993; Bossard et al., 1996). ECM degradation is also believed to play an important role in ovarian function. The differentiation of germinal cells into primordial follicles, their further growth into functional ovulatory follicles and their rupture, plus the formation of corpus luteum and its subsequent atresia, all involve cell migration and displacement, and the destruction and repair of ECM (Woessner, 1982, 1991). Accordingly, we have recently shown that the mRNA levels for structural components of the ovarian ECM exhibit >2-fold changes over the 4-day oestrous cycle of the mouse (Oksjoki et al., 1999). Similar oestrous cycle-dependent changes have also been demonstrated in the production and activity of MMPs and their inhibitors (tissue inhibitors of matrix metalloproteinases, TIMPs) during follicular development (Hulboy et al., 1997; Duncan et al., 1998; Oksjoki et al., 1999). However, essentially nothing is known about the involvement of cathepsins in ECM degradation and other proteolytic processes in the ovary. Cysteine cathepsins have been shown to participate in the activation of prorenin to renin in kidneys and salivary glands (Morris, 1992; Sano et al., 1993), and in the stimulation of steroidogenesis in the testis (Boujrad et al., 1995), but it is not known whether the same situation exists in the ovary.
Considering this background and the availability of cDNA clones and antibodies to the major cysteine cathepsins, it is surprising that no systematic analyses are available on their production and distribution in the ovary either in the literature or in the ovarian kaleidoscope database (http://ovary.stanford.edu). In the present study we attempt to fill this gap, and report on the mRNA levels of cathepsins B, H, K, L and S at different stages of the 4-day oestrous cycle of the mouse and on the cellular distribution of the mRNAs and proteins during follicular development.
Materials and methods
Ovarian samples
This study is based on the analysis of ovaries from 45 C57 blxDBA mice which were housed in a pathogen-free animal facility under controlled lighting conditions with lights on from 05.00 to 19.00 h. The animals were given water and pelletted food ad libitum. Vaginal cytology was used for determination of the oestrus cycle, which was divided into six phases (di-oestrus, early pro-oestrus, late pro-oestrus, oestrus, meta-oestrus I, meta-oestrus II) according to vaginal cell morphology after Papanicolau staining as described in detail earlier (Oksjoki et al., 1999). To obtain representative samples, the oestrous cycle of each animal was followed by analysis of the vaginal smears at four consequtive days before the animal was killed by cervical dislocation. The ovaries were excised, trimmed free from surrounding tissues, and frozen at –80°C for RNA extraction or fixed in 4% paraformaldehyde, embedded in paraffin and sectioned serially into 5 µm sections. The study protocol was approved by the institutional committee for animal welfare.
RNA extraction and mRNA analyses
For extraction of total RNA, the frozen ovaries were pulverised under liquid nitrogen in a mortar and dissolved in guanidinium isothiocyanate as described previously (Chirgwin et al., 1979). Aliquots (10 µg) of total RNA were denatured with glyoxal and formamide, fractionated on 0.75% agarose gels, and blotted onto nylon transfer membranes, and hybridized with [32P]-labelled cDNA inserts at 42°C for 20 h. The hybridizations and washes were performed as suggested by the supplier (Pall BioSupport Division, Glen Cove, NY, USA). Inserts of cDNA clones pMCatB-1, pMCatH-1, pMCatL-1, pMCatS-1 (Söderström et al., 1999) and pMCatK-2 (Rantakokko et al., 1996) were used as probes for mouse cathepsin B, H, L, S and K mRNAs respectively. The bound probes were detected and quantified on a Molecular Imager phosphoimager and the signals corrected for variations in the 28S rRNA levels determined by hybridization.
Immunohistochemistry
Formalin-fixed, paraffin-embedded histological sections of normal mouse ovaries were deparaffinized, rehydrated and digested for 1 h with hyaluronidase (2 mg/ml) in phosphate-buffered saline (PBS, pH 5). Immunohistochemistry for cathepsins B, H, K, L and S was performed using polyclonal antibodies described earlier (Söderström et al., 1999). The bound antibodies were detected as brown precipitate using the avidin–biotin complex method (Vecstatin ABC kit; Vector, Burlingame, CA, USA). The sections were counterstained with haematoxylin. The specificity of the immunoreactions was controlled by omitting the primary or secondary antibody, and by replacing the primary antibody with preimmune serum. The intensity of immunostaining at the different stages of follicular development was evaluated independently by each author.
In-situ hybridization
Formalin-fixed, paraffin-embedded histological sections were processed for in-situ hybridization as described earlier (Sandberg and Vuorio 1987, Ylä-Herttuala et al., 1990). The sections were deparaffinized, treated with 0.2 mol/l HCl and digested with proteinase K (5 µg/ml) in PBS. The hybridizations were performed using both antisense and sense cRNA probes synthesized by T7 and SP6 RNA polymerases using linearized plasmids as templates and [35S]-UTP as the labelled nucleotide. After ribonuclease treatment and washes, the bound probes were detected autoradiographically. The sections were counterstained with haematoxylin.
Results
Northern blot analyses
Determination of mRNA levels of cathepsins B, H, K, L and S at specific stages of the oestrus cycle revealed some consistent changes (Figure 1). Although the expression profiles for individual cathepsin mRNAs varied somewhat, the highest mRNA levels for each cathepsin were observed in ovaries at late pro-oestrus and oestrus (Figure 2).
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Immunohistochemical localization of cathepsins
In contrast to the relatively small degree of cyclic variation in the mRNA levels of the cathepsins studied, striking differences were observed in the immunohistochemical localization of the corresponding proteins through follicular development and growth, and during corpus luteum formation (Figure 3
). As summarized in Table I, each cathepsin revealed a unique distribution pattern within the ovary.
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Cathepsin B was predominantly seen in the germinal epithelium and the underlying cell layers throughout the oestrous cycle (Figure 3A1-4
). Another typical feature of the germinal epithelium was its patchy immunostaining for cathepsin H, and to some extent also for cathepsins K and S, but lack of staining for cathepsin L (Figure 3). Intense staining for cathepsins B, H and K was typically seen in germinal epithelium in contact with secondary (Figure 3) and Graafian follicles (Figure 3).
A characteristic feature of the theca cell layer was its intense staining for cathepsin H. This staining was sharply defined to one or two cell layers in theca externa (Figures 3B and 4B). This cellular staining pattern became discontinuous upon follicular involution (Figure 5A) and largely disappeared during corpus luteum formation (Figure 3B). A more diffuse and weaker staining of the theca cell layer was also seen with antibodies specific for cathepsin K (Figure 4C) and S (Figure 4E).
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Follicular development was accompanied by an increasing number of granulosa cells becoming immunopositive for cathepsins H (Figure 4B
) and S (Figure 4E). Upon follicular maturation individual granulosa cells also became immunopositive for cathepsins B, K and L. The oocyte exhibited immunopositivity for each cathepsin studied (Figures 3 and 4).
Corpus luteum was the predominant location of cathepsin L immunostaining with the highest levels in the large cuboidal cells of small, developing corpora lutea (Figures 3D5, 4D and 5B). Such cells also stained positive for cathepsin S (Figure 3E5), whereas the weaker staining for cathepsins H and K (Figure 3B5 and 3C5 respectively) may only reflect the tendency of the secondary antibody to non-specifically bind to corpus luteum cells (Figure 3F5). The immunostaining of corpora lutea for cathepsins L and S gradually decreased upon ageing, in association with the size reduction and morphological change of the luteal cells (Figure 5B).
The connective tissue stroma of the mouse ovary exhibited essentially no staining for cathepsins B, H, K, L and S (Figure 3), whereas control stainings with preimmune sera showed a reaction with stromal cells to some extent (Figure 4F).
Cellular localization of cathepsin mRNAs
Finally, in-situ hybridization of serial sections used for immunohistochemistry was performed to localize the mRNAs coding for cathepsins B, H, K, L and S. The distribution of the mRNAs in some, but not all, granulosa cells (Figure 6) agreed well with the immunohistochemical localization of the corresponding proteins (Figure 4). Similarly, the mRNA for cathepsin L was enriched in the cells of developing corpora lutea (Figure 6E), and the mRNA for cathepsin B in germinal epithelium and underlying cell layers (Figure 6A). The enrichment of mRNAs for cathepsins H (Figure 6C), K and S (Figure 6D) in theca and granulosa cells was not as obvious as would be expected from the immunohistochemical distribution of the proteins.
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Discussion
The present study demonstrates complex temporal and spatial expression patterns of cathepsins B, H, K, L and S in the mouse ovary at different stages of follicular development. We have also demonstrated expression of cathepsin mRNAs in the whole ovary throughout the 4-day oestrous cycle, although direct comparison with protein localization during follicular development cannot be made. Even though changes observed in cathepsin mRNA levels during the oestrous cycle were smaller than those observed in the mRNA levels for structural components of the ECM (Oksjoki et al., 1999), the differential distribution of the specific cathepsins and their mRNAs in the mouse ovary suggests that these enzymes participate in different proteolytic processes. Similar oestrous cycle-dependent changes have been demonstrated in the production and activity of MMPs and TIMPs during follicular development and luteal rescue (Hulboy et al., 1997; Duncan et al., 1998; Oksjoki et al., 1999). No earlier data are available on the stage-specific expression patterns of cathepsin mRNAs in the ovary, although their presence in unspecified ovary RNA samples has been reported previously (Petanceska and Devi, 1992; Brömme and Okamoto, 1995; Rantakokko et al., 1996; Kirschke et al., 1998; Söderström et al., 1999). Recently, induction of cathepsin L production into follicular fluid of preovulatory follicles was reported during analysis of transgenic mice null for the progesterone receptor gene (Robker et al., 2000). These observations are in agreement with our finding of cathepsin L and its mRNA in granulosa cells during follicular development.
Based on the results of the present study, several different functions can be proposed for specific cathepsins in the ovary. The predominant expression of cathepsin B in the germinal epithelium, particularly at sites where the Graafian follicle is in contact with the germinal epithelium, suggests that this enzyme is involved in follicular rupture. This function could be both direct and indirect, as cathepsin B has also been shown to stimulate the activation of MMPs (Kostoulas et al., 1999). This stimulation occurs indirectly by proteolytic degradation of TIMPs without an increase in the transcription of MMP genes. The patchy localization of cathepsins H, K and S in germinal epithelium overlying the secondary and Graafian follicles is also consistent with increased proteolysis at sites where thinning of the collagenous layers surrounding the follicle occurs prior to ovulation (Espey, 1967). Increases have also been observed in the mRNA levels for MMP-1 and MMP-2 (Reich et al., 1985, 1991; Hulboy et al., 1997) as well as for TIMP-1 and TIMP-3 (Nothnick et al., 1995; Inderdeo et al., 1996) at the time of ovulation.
The increased presence of cathepsins H and S in some, but not all, granulosa cells suggest these enzymes play some role in follicular maturation. Gradually, individual granulosa cells also stained positive for the other cathepsins studied. These staining patterns suggest heterogeneity of the granulosa cell population in the mouse ovary. In some cases, the cathepsin-positive cells surrounded the maturing oocyte which itself stained positive for all the cathepsins studied.
The presence of cathepsins in theca cells is consistent with the active proteolysis which must accompany the increase in size of the follicle. Cathepsin H exhibited the strongest staining with a particularly well-defined distribution limited to one or two layers of theca cells and is, therefore, likely to play a role in the fragmentation of the theca basement membrane. When the follicle reached the Graafian stage, this cell layer became discontinuous, but cathepsin H immunostaining persisted in the fragmented basement membrane even after follicular rupture (Figures 3B5 and 5A). Cathepsins K and S were also found in the theca cell layer, but their distribution covered all theca cells, including the inner layer involved in steroid production (Azziz et al., 1997).
During formation of the corpus luteum, the disruption of the basement membrane was associated with gradual disruption of the theca cell layer, disappearance of their staining for cathepsins H, K and S, and the appearance of strong immunostaining for cathepsin L in the luteal cells. This observation of high levels of cathepsin L and its mRNA in active corpora lutea is consistent with the suggested role of a complex of procathepsin L and TIMP-1 in stimulation of steroidogenesis (Boujrad et al., 1995), as the corpus lutem is a site of very active steroid production. The mRNA levels of TIMP-1 in the mouse have also been shown to increase during luteolysis (Inderdeo et al., 1996), and the presence of the corresponding protein predominantly in large luteal cells has been reported (Smith et al., 1996). The formation of corpus luteum has been compared with the connective tissue repair process, both by histological appearance (Woessner, 1982), and by the ratio of type III and I collagen mRNAs typical for granulation tissue (Oksjoki et al., 1999). The lack of cathepsin immunostaining in the connective tissue stroma of the ovary suggests that matrix degradation primarily results from the activity of granulosa, germinal epithelial and luteal cells.
In addition to lysosomal and extracellular degradation of proteins, and stimulation of steroidogenesis (Kirschke et al., 1998), suggested biological functions of cathepsins in the ovary include activation of prorenin to renin in the kidney (Morris, 1992; Neves et al., 1996). In the human, the ovary is an important site for conversion of prorenin to renin, as indicated by abnormal processing associated with polycystic ovary syndrome (PCOS) (Jaatinen et al., 1995). On the other hand, the changes in cathepsin mRNA levels in late pro-oestrus, coinciding with the LH surge (Rugh, 1990), could indicate that their expression is regulated by gonadotrophins.
Despite demonstration of the differential distribution of specific cathepsins in the mouse ovary, their roles in ovarian function remain hypothetical. Detailed studies on transgenic mice over-expressing specific cathepsin genes and knock-out mice with inactivating mutations of these genes, should provide important data on the specific roles of cysteine cathepsins in ovarian physiology. Additional data on the putative functions of the different cathepsins can also be obtained by comparing the cellular localization of metabolic processes and expression patterns of other genes in the ovary. For such comparisons the value of the gene expression database (http://ovary.stanford.edu) containing compiled data on the temporo spatial expression patterns of an increasing number genes is obvious. Our observations also warrant further studies on the spatial and temporal distribution of cathepsin mRNAs in human ovaries, not only during the normal menstrual cycle, but in diseases affecting follicular maturation, e.g. PCOS, for a better understanding of the underlying metabolic aberrations.
Acknowledgments
The authors are grateful to Päivi Auho, Tuula Oivanen and Anu Kupiainen for expert technical assistance. Drs Heidrun Kirschke and Dieter Brömme are acknowledged for providing the cathepsin antibodies. This study was financially supported by grants from the Academy of Finland (project no 37311), Sigrid Juselius Foundation and the Turku University Central Hospital (project no 13449). Sanna Oksjoki is a recipient of a training grant from Turku Graduate School of Biomedical Sciences.
Notes
3 To whom correspondence should be addressed at: Family Federation of Finland, Infertility Clinic of Turku, Maariankatu 3a, FIN-20100 Turku, Finland. E-mail: marja-leena.anttila{at}vaestoliitto.fi 
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V. Jokimaa, S. Oksjoki, H. Kujari, E. Vuorio, and L. Anttila Expression patterns of cathepsins B, H, K, L and S in the human endometrium Mol. Hum. Reprod., January 1, 2001; 7(1): 73 - 78. [Abstract] [Full Text] [PDF]
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