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BOR - Papers in Press, published online ahead of print August 20, 2003.
Biol Reprod 2003, 10.1095/biolreprod.103.020826
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BIOLOGY OF REPRODUCTION 69, 1998–2006 (2003)
DOI: 10.1095/biolreprod.103.020826
© 2003 by the Society for the Study of Reproduction, Inc.

Ovary

Spatial Expression Patterns of Activin and Its Signaling System in the Zebrafish Ovarian Follicle: Evidence for Paracrine Action of Activin on the Oocytes1

Yajun Wang, and Wei Ge2

Department of Biology, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
We have previously demonstrated that activin is likely an ovarian mediator of pituitary gonadotropin(s) and local epidermal growth factor in their stimulating oocyte maturation and maturational competence in the zebrafish. However, the downstream events controlled by activin remain unknown. One possible mechanism is that activin may directly work on the oocytes to promote the development of oocyte maturational competence. To substantiate this hypothesis, we performed the present study to demonstrate the expression of the activin system in different compartments of zebrafish follicles, namely, the follicle cells and oocytes. The proteins examined include activin subunits (ßA and ßB), activin-binding protein (follistatin), activin type II receptors (type IIA and IIB), the type I activin receptor-like kinases (ALK1-like, ALK2-like, and ALK4-like), and the intracellular activin signaling molecules (Smad2, Smad3, Smad4, and Smad7). The results showed that the entire activin signaling system is expressed by the full-grown immature zebrafish oocytes (~0.65 mm in diameter), including ALK4-like (ActRIB), ALK2-like (ActRIA), ActRIIA, ActRIIB, Smad2, Smad3, Smad4, and Smad7, therefore supporting our hypothesis that the oocytes are one of the direct targets of activin actions in the zebrafish ovary. In contrast, activin itself (ßA and ßB) and ALK1-like type I receptor are predominantly expressed in the follicle cells surrounding the oocytes. Interestingly, although follistatin is expressed in both the follicle cells and oocytes, its level of expression is significantly higher in the oocytes than the follicle cells, implying that follistatin may serve as a signal from the oocytes to modulate the activity of activin produced by the follicle cells. Taken together, the present study provides convincing evidence that although all members of the activin system are expressed in the whole follicle, they exhibit distinct spatial patterns of expression among different compartments of the follicle. It is likely that activin works directly on the oocytes in a paracrine manner to promote oocyte maturation and maturational competence. On the other hand, instead of being controlled passively by the follicle cells, the oocytes may actively participate in the regulation of follicle development by releasing various modulating molecules such as follistatin.

activin, follicle, follistatin, ovary


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Activin was originally identified as a peptide growth factor from the ovarian follicular fluid based on its stimulation of FSH secretion in cultured anterior pituitary cells [1, 2]. It is a homodimer or heterodimer of two similar but distinct ß subunits (ßA and ßB). The dimerization of activin ß subunits gives rise to three forms of activin: activin A (ßAßA), activin B (ßBßB), and activin AB (ßAßB) [3]. The activities of activin are modulated by its binding protein follistatin [4], which was originally purified from mammalian follicular fluid as an FSH inhibitor [5–7]. Follistatin binds activin with high affinity, and its binding effectively neutralizes the bioactivities of activin in a variety of target tissues [4, 8].

Activin acts through specific receptors that belong to a family of membrane serine/threonine kinases. Activin signaling involves two types of transmembrane receptors, namely, activin type I and type II receptors. There are two subtypes of activin type II receptors in most species studied, namely, type IIA (ActRIIA) and type IIB (ActRIIB), respectively [9–11]. The type I receptors refer to a family of closely related activin receptor-like kinases (ALK) that transduce signals for different members of the transforming growth factor ß (TGFß) superfamily. Among all ALKs studied so far, ALK2 (ActRIA) and ALK4 (ActRIB) have been documented to be involved in activin signaling. Activin acts by first binding to a type II receptor, which in turn recruits and activates a type I receptor by phosphorylation [10–12]. The activated complex of activin and its receptors then stimulate the downstream intracellular signaling molecule (Smad) proteins. Three types of Smads are involved in activin signaling: the receptor-activated Smads (Smad2 and Smad3), the common Smads (Smad4), and the inhibitory Smads (Smad7). The binding of the receptors by activin directly activates Smad2 or Smad3, which in turn associates with the common Smad4. The Smad complex then translocates into the nucleus and regulates target gene transcription with cofactors [13–15].

Activin has diverse activities in the ovary of mammals. Recombinant activin stimulates follicle growth [16] and granulosa cell proliferation [17] and increases FSH receptor and FSH-induced LH receptor production [18–20]. Activin also regulates ovarian steroidogenesis, including stimulating estradiol release by the granulosa cells [21–23] but suppressing androgen production in the theca cells [24]. In addition, several lines of evidence in mammals have implicated activin in promoting oocyte maturation [25–28] and developmental competence [29, 30]. However, there are also reports that activin has no effect on oocyte maturation in the rat [31, 32] and pig [33]. Therefore, the issue of paracrine regulation of oocyte development by activin requires more studies in different models and under different experimental conditions.

The production of activin ßA and ßB in the ovary has been localized mainly in the granulosa cells of mammalian ovarian follicles by in situ hybridization or immunohistochemistry [34–37]. Similarly, activin receptors [38–41] and their downstream signaling Smad proteins [42, 43] have also been demonstrated in the follicle cells. Interestingly, follistatin has also been localized predominantly in the granulosa cells in several species, including the rat [44], sheep [45], and human [36], by in situ hybridization. Follistatin effectively antagonizes the actions of activin in the ovary, and its expression in the ovary suggests that the activities of ovarian activin are likely subject to the modulation by local follistatin [4].

Compared with the extensive studies on activin in mammalian ovaries, the roles of activin in the ovary of lower vertebrates, such as fish, remain relatively unknown. The expression of activin subunits has been immunocytochemically localized to the follicle cells and previtellogenic oocytes in the goldfish, medaka, and rainbow trout ovary [46–48]. Using zebrafish as the experimental model, we have recently demonstrated that activin stimulates zebrafish oocyte maturation [49] and strongly promotes the development of oocyte maturational competence in vitro [50] and that the ovarian activin system is likely one of the downstream mediators of pituitary gonadotropin(s) and local epidermal growth factor (EGF) in the zebrafish ovary [49, 51]. However, what exactly activin does in the zebrafish ovary remains largely unknown. Activin does not seem to promote final oocyte maturation by increasing the secretion of 17{alpha},20ß-dihydroxy-4-pregnen-3-one (DHP), the most potent maturation-inducing steroid in most teleosts studied, because it does not have significant effect on the expression of 20ß-hydroxysteroid dehydrogenase [52] and 17{alpha}-hydroxylase/17,20-lyase (unpublished results), the two steroidogenic enzymes critical for the synthesis of DHP.

Considering that activin has a potent effect on the development of oocyte maturational competence in the zebrafish, which is comparable to that of gonadotropin [50], we hypothesize that activin may enhance the rate of final oocyte maturation by directly acting on the oocytes to enhance their maturational competence or responsiveness to DHP. To substantiate our hypothesis, we must provide evidence for the existence of activin signaling system in the oocytes, including its receptors and postreceptor signaling molecules. The present study was therefore undertaken to investigate the spatial expression patterns of activin ßA and ßB subunits, follistatin, activin receptors (ActRIIA, ActRIIB, and ALKs), and Smads (Smad2, Smad3, Smad4, and Smad7) in the ovarian follicles of the zebrafish using semiquantitative reverse transcriptase-polymerase chain reaction (RT-PCR).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

Zebrafish, Danio rerio, were purchased from local pet stores and maintained in the flow-through aquaria (36L) at 26°C on a 14L:10D photoperiod. The fish were fed twice a day with commercial tropical fish food with supplement of live brine shrimp once or twice a week. All experiments were performed under license from the Government of the Hong Kong Special Administrative Region of the People's Republic of China and endorsed by the Animal Experimentation Ethics Committee of the Chinese University of Hong Kong.

Preparation of Total RNA from Oocytes, Follicle Layers, and Cultured Follicle Cells

Ovaries were removed from 15 to 20 sexually mature female zebrafish after decapitation and placed in a 90-mm culture dish containing 60% Leibovitz L-15 medium (Gibco Invitrogen, Carlsbad, CA). The full-grown immature follicles (~0.65 mm) were carefully separated (Fig. 1). Sixty healthy follicles were selected and transferred into a 1.5-ml tube with 400 µl of medium at 4°C. To isolate total RNA from the oocytes, the follicles were gently pressed with a homogenizer pestle once to release the ooplasm. After centrifugation at 3000 rpm briefly for 30 sec, 200 µl of the supernatant was quickly transferred into a new 1.5-ml tube containing 1.0 ml of Tri-Reagent (Molecular Research Center, Cincinnati, OH) at 4°C to extract total RNA according to the manufacturer's protocol. The pellet of the follicle layers was pressed one more time with a new pestle to ensure that all follicles were ruptured with minimal amounts of ooplasm enclosed. The follicle layers were then vigorously washed three times with 1 ml of L-15 medium at 4°C to minimize ooplasm contamination. Total RNA was then extracted from the follicle layers with Tri-Reagent.



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  		FIG. 1. A) Cross section of a full-grown immature zebrafish follicle (x40). B) A close-up of the follicle layer (arrow) on the surface of the oocyte (x400). GV, germinal vesicle

To confirm that the expression of genes in the follicle layers is not due to ooplasm contamination, we also analyzed their expression in the primary culture of zebrafish follicle cells that is free of oocytes. The follicle cells were cultured as described previously [53, 54].

Reverse Transcription-Polymerase Chain Reaction

RT was performed at 42°C for 2 h in a total volume of 10 µl consisting of 3 µg of RNA, 1x Single Strand Buffer (Gibco Invitrogen), 10 mM dithiothreitol, 0.5 mM each deoxyribonucleotide triphosphate (dNTP), 0.5 µg of oligo(dT), and 100 U of SuperScript II (Gibco Invitrogen). One microliter of the reaction mixture was subjected to PCR amplification using gene-specific primers (Table 1). The amplification reaction was performed in a volume of 30 µl consisting of 1x PCR buffer, 0.2 mM each dNTP, 2.5 mM MgCl2, 0.2 µM of each primer, and 0.6 U of Taq polymerase on the Thermal Cycler 9600 (Eppendorf, Hamburg, Germany). To detect the expression of genes in the cultured follicle cells, 32 cycles of reaction were performed with profiles of 30 sec at 94°C, 30 sec at 56°C (follistatin, activin ßA, and activin ßB) or 58°C (other genes) and 60 sec at 72°C.


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  		TABLE 1. Primers used in the RT-PCR assays.a

Validation of Semiquantitative RT-PCR Assays

To optimize the cycle numbers used for semiquantitative PCR analysis, RT-PCR reactions were performed as described above using RNA isolated from the immature full-grown follicles. PCR was performed in a volume of 30 µl for various cycles, and the products from different cycles of amplification were visualized on a UV-transilluminator after electrophoresis on 1.8% agarose gel containing ethidium bromide. The signal intensity was quantitated with the Gel-Doc 1000 system and Molecular Analyst Software (Bio-Rad, Hercules, CA). The cycle numbers that generate half-maximal amplification were used for subsequent semiquantitative analysis of gene expression, and they are 27 cycles for ALK4-like receptor, Smad2, and Smad3, 28 cycles for ALK1-like and ALK2-like receptors, 32 cycles for ActRIIB, Smad4, and Smad7, and 21 cycles for ß-actin. The validation of RT-PCR assays for activin ßA, activin ßB, ActRIIA, and follistatin had been reported in our previous studies, and the cycle numbers used for these genes are 29–31 cycles for activin ßA, activin ßB, and follistatin [54, 55] and 29 cycles for ActRIIA [53]. The specificity of PCR amplification was confirmed by either sequencing the PCR products directly or cloning the PCR products into pBluescript II KS (+) (Stratagene, La Jolla, CA) followed by sequencing. To further validate the semiquantitative RT-PCR assays, PCR amplification (30 µl) was performed on 3 µl of serially diluted purified PCR products of these genes using the cycle numbers optimized above to evaluate the correlation between the input of template and the output of PCR amplification (Fig. 2).



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  		FIG. 2. Validation of the semiquantitative RT-PCR assays. PCR amplification of serially diluted templates of ActRIIB, ALK4-like, ALK2-like, ALK1-like receptor, Smad2, Smad3, Smad4, and Smad7 was performed using the cycle number that generates half maximal reaction for each gene to evaluate the correlation between the template amount and PCR output. Each value represents the mean ± SEM of three PCR reactions. The gel images of the PCR products are shown at the lower right corner with the triplicate reactions on each diluted template marked by the labels on the top

Data Analysis

The mRNA level of each gene was first calculated as the ratio to that of ß-actin, which was amplified as the internal control and then expressed as the percentage of the mRNA level in the follicle layers. The data were analyzed by the Student t-test using GraphPad Prism 3.0cx for Macintosh OS X 10.2 (GraphPad Software, San Diego, CA). The experiment was repeated twice, and independent RNA isolation, RT reaction, and PCR were performed in quadruplicate in each experiment.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Evidence for Clean Separation of RNA from the Oocytes and Follicle Layers

Although our method of separating oocytes and follicle layers is efficient, minimal contamination of the ooplasm by the follicle cells and vice versa cannot be avoided. To demonstrate that the expression of genes detected in the isolated follicle layers is not due to contamination by the ooplasm, we first examined their expression in the primary culture of zebrafish follicle cells, which is free of oocytes. The results showed that all genes examined were expressed in the cultured follicle cells. No signal was detected in the RNA without reverse transcription (Fig. 3).



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  		FIG. 3. Expression of activin subunits (ActßA and ActßB), follistatin (FS), activin type II receptors (ActRIIA and ActRIIB), type I-like receptors (ALK4-like, ALK2-like, and ALK1-like), and the Smads (Smad2, Smad3, Smad4, and Smad7) in cultured zebrafish ovarian follicle cells. +, with reverse transcription; -, RNA without reverse transcription

Since FSH and LH receptors are expressed only in the follicle cells [56–58], we used the transcripts of these two genes as the markers to evaluate the contamination of the oocyte RNA preparation by that from the follicle layers. As expected, strong signals for both FSH and LH receptors were detected in the RNA from the follicle layers, whereas only very faint signals were detected in that of the oocytes, indicating that the total RNA prepared from the oocytes was fairly clean, with little contamination from the follicle cells, and therefore suitable for quantitating the relative abundance of gene expression in these two follicle compartments (Fig. 4).



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  		FIG. 4. Expression of FSH receptor (FSHR) and LH receptor (LHR) in isolated follicle layers and oocytes. The expression levels are normalized by ß-actin and expressed as the percentage of the mRNA level in the follicle layer. Each value represents the mean ± SEM of four independent RT-PCR reactions. ***P < 0.001 vs. follicle layer

Relative Abundance of Activin ßA, Activin ßB, and Follistatin mRNA in the Zebrafish Oocytes and Follicle Layers

Our previous studies suggest that the ovarian activin system is likely involved in the actions of pituitary gonadotropin and local EGF on oocyte maturation [49–51]. To determine the source of production of activin subunits (ßA and ßB) and its binding to protein follistatin in the follicle, we evaluated their relative levels of expression in the zebrafish oocytes and follicle cells.

The RT-PCR assays for activin ßA, activin ßB, and follistatin had been carefully validated in our previous studies [54, 55]. Activin ßA and ßB subunits were predominantly expressed in the follicle layers (Fig. 5). Although signals for activin subunits, especially activin ßA, could also be detected in the oocytes, they were significantly lower than those in the follicle cells. Despite the fact that the oocytes may express activin as demonstrated previously in both fish [46] and mammals [41], minor contamination of RNA from the follicle cells might also contribute to the signals detected. Interestingly, the expression pattern of the activin-binding protein follistatin was opposite to that of activin subunits. Its expression level was significantly higher in the oocytes than that in the follicle layers (Fig. 5).



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  		FIG. 5. Expression of activin ßA, activin ßB, and follistatin in isolated follicle layers and oocytes. The expression levels are normalized by ß-actin and expressed as the percentage of the mRNA level in the follicle layer. Each value represents the mean ± SEM of four independent RT-PCR reactions. ***P < 0.001 vs. follicle layer

Evidence for the Expression of Activin Receptors and Smads in the Zebrafish Oocytes

Although there is no doubt that the ovarian follicle cells are both the sites of activin production and the targets of its actions because its receptors and Smads are all expressed in these cells (Fig. 3), our previous evidence implied that activin may also act directly on the oocytes to mediate the effect of pituitary gonadotropin on oocyte maturational competence [49]. To provide evidence for this hypothesis, we analyzed the expression of activin signaling system in the oocytes compared with their expression levels in the follicle layers. The molecules examined included activin type II receptors (ActRIIA and ActRIIB), three cloned zebrafish ALKs (Acvrl1 [59], zALK8 [60], and TARAM-A [61]), and Smads (Smad2, Smad3, Smad4, and Smad7). The ligand identity of the three type I receptors (Acvrl1, zALK8, and TARAM-A) remains unknown; however, sequence analysis showed that they are closely related to mammalian ALK1, ALK2 (ActRIA), and ALK4 (ActRIB), respectively (Fig. 7). ALK2 and ALK4 have been well documented in activin signaling, whereas ALK1 is likely involved in TGFß or bone morphogenetic protein signaling [12, 14, 62, 63].



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  		FIG. 7. Phylogenetic analysis of the ALK gene family. The tree based on the full amino acid sequences of major ALKs was constructed by neighbor-joining method [74] with bootstrap analysis (1000 replicates) in MEGA 2. Numbers at the nodes indicate bootstrap values in percentage. The length of horizontal lines shows the genetic distance estimated using Kimura's two-parameter substitution model [75], with reference to the horizontal scale bar. The length of vertical lines is arbitrary and for clarity only

ActRIIA and ActRIIB were expressed in both the oocytes and follicle layers. However, unlike ActRIIB, which was equally expressed in both compartments, the level of ActRIIA was slightly but significantly higher in the oocytes (Fig. 6). As for the expression of type I receptors, the expression levels of zALK8 (ALK2-like) and TARAM-A (ALK4-like) in the oocytes were both significantly higher than those in the follicle layers. In contrast, the expression of Acvrl1 (ALK1-like) was exclusively expressed in the follicle layer with little signal detected in the oocytes (Fig. 8). All the Smad proteins examined were expressed in both the oocytes and follicle layers. Smad2 and Smad7 were equally expressed in the oocytes and follicle layers, whereas the expression of Smad3 and Smad4 was significantly higher in the oocytes compared with that in the follicle layers (Fig. 9).



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  		FIG. 6. Expression of activin type IIA and IIB receptors in isolated follicle layers and oocytes. The expression levels are normalized by ß-actin and expressed as the percentage of the mRNA level in the follicle layer. Each value represents the mean ± SEM of four independent RT-PCR reactions. *P < 0.05 vs. follicle layer



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  		FIG. 8. Expression of zebrafish ALK4 (ActRIB)-like, ALK2 (ActRIA)-like, and ALK1-like type I receptors in isolated follicle layers and oocytes. The expression levels are normalized by ß-actin and expressed as the percentage of the mRNA level in the follicle layer. Each value represents the mean ± SEM of four independent RT-PCR reactions. *P < 0.05; **P < 0.01; ***P < 0.001 vs. follicle layer



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  		FIG. 9. Expression of Smad2, Smad3, Smad4, and Smad7 in isolated follicle layers and oocytes. The expression levels are normalized by ß-actin and expressed as the percentage of the mRNA level in the follicle layer. Each value represents the mean ± SEM of four independent RT-PCR reactions. *P < 0.05; **P < 0.01 vs. follicle layer


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Both activin ßA and ßB are expressed in the zebrafish ovary; however, the types of cells in the full-grown follicles that express these proteins remain unknown [49, 51, 53, 54, 64]. In the present study, we demonstrated that activin ßA and ßB are predominantly expressed in the follicle layers of the full-grown follicles, which agrees well with our earlier report in the goldfish that strong immunocytochemical staining for activin ßA and ßB could be observed in the follicle cells of vitellogenic and full-grown follicles [46] and similar studies in the rainbow trout ovary [47]. Our results are also consistent with those in mammals and humans that the ovarian granulosa cells are the major expression sites of both activin ß subunits [34–37, 39]. What remains interesting and controversial is whether the oocytes also produce activin molecules. In the human and mouse, no ßA mRNA was detectable in the oocytes by RT-PCR [39]. Similar results have also been reported in Xenopus laevis [65]. However, in the rat and cow, ßA mRNA could be consistently detected in the oocytes by RT-PCR and immunostaining [26, 41, 66, 67]. The discrepancy among these studies could probably be due to different species or stage of oocytes used. The stage-dependent expression of activin in the oocytes has been observed in the goldfish and medaka, with the immunoreactive activin subunits (ßA and ßB) being detected in the previtellogenic oocytes but not the later stages [46, 48]. Although we cannot rule out the possibility that the full-grown oocytes also produce activin, the present study showed that they are not the major production sites compared with the follicle cells at this stage.

One of the interesting findings of the present study is that the expression of follistatin is significantly higher in the oocytes than in the follicle cells, in contrast to that of activin subunits. This is similar to the situation reported in the cow, whose oocytes exhibited strong immunoreactivity for follistatin [41], but different from that in other mammalian species, including humans. In the rat, sheep, and human, follistatin is abundantly expressed in the granulosa cells as demonstrated by in situ hybridization [36, 44, 45], and only weak signals were detected in the human oocytes when compared with that in the cumulus cells [39]. No follistatin mRNA was detected in the mouse oocytes by RT-PCR [39]. Since follistatin is a secreted protein that binds and neutralizes activin [4], its high expression by the zebrafish oocytes implies that the oocytes may actively participate in the development of follicles by releasing follistatin to modulate the activities of activin secreted by the follicle cells. Further studies on the temporal pattern of follistatin expression in the developing oocytes and its regulation will provide clues to its roles in follicle development.

We have previously demonstrated that both human activin A and goldfish activin B are potent in promoting zebrafish oocyte maturational competence, the responsiveness to DHP [50]. One of the possible mechanisms underlying the action of activin is that it may directly act on the oocytes to stimulate the biosynthesis of DHP receptors, which appear concurrently with the development of oocyte maturational competence [68, 69]. To provide support for this hypothesis, we further examined the expression of key components of activin signaling system in the zebrafish oocytes, which would provide compelling evidence for direct activin action on the oocytes. The results showed that the entire activin signaling system, including activin type II receptors (ActRIIA and ActRIIB), ALK2- and ALK4-like type I receptors, and Smad proteins known to be involved in the activin signaling pathway (Smad2, Smad3, Smad4, and Smad7), was expressed by the oocytes. Moreover, most of these proteins exhibited higher expression levels in the oocytes than in the follicle layers, further supporting the idea that the zebrafish oocytes are potential targets for direct activin actions.

The demonstration of ActRIIB expression in the zebrafish oocytes agrees with a previous report using in situ hybridization that showed that ActRIIB could be detected in the zebrafish oocytes of different stages [70]. The expression of ActRIIA and ActRIIB in the oocytes has also been demonstrated in a variety of mammalian species using RT-PCR, in situ hybridization, or immunohistochemical staining [26, 38–43, 66, 67], suggesting that a paracrine influence on the oocytes by activin from the follicle cells is likely a well-conserved regulatory mechanism in vertebrate follicles. In contrast to the consistent detection of activin type II receptors in vertebrate oocytes, the expression of activin type I receptors, ActRIA (ALK2) and ActRIB (ALK4), in the oocytes varies among different species or stages of development in mammals [39, 71]. In the human and mouse, both ActRIA and ActRIB are expressed in the oocytes, whereas in the rat, ActRIA was mainly expressed in the oocytes of preantral and antral follicles and ActRIB was not detectable by in situ hybridization [71]. In immature bovine oocytes, little expression of ActRIB was detected by RT-PCR [67]. In the present study, our results clearly showed that the ALK2- and ALK4-like receptors were both abundantly expressed in the zebrafish oocytes. In contrast, the expression of ALK1-like receptor was exclusively restricted to the follicle layers.

It is generally believed that the receptor-activated Smad2 and Smad3, the common Smad4, and the inhibitory Smad7 are responsible for activin signaling within cells [14, 72]. To provide further evidence that the oocytes are the direct targets of activin in the zebrafish, we also examined the expression of these Smads in the oocytes. All Smad proteins studied, Smad2, Smad3, Smad4, and Smad7, were expressed in the zebrafish oocytes, and their expression levels were either equal to or higher than those in the follicle layers. This agrees with recent studies in the rat and human using immunostaining and RT-PCR [42, 43, 73]. Although both Smad2 and Smad3 have been demonstrated in activin signaling [15], their involvement in activin signal transduction in the zebrafish oocytes needs to be further elucidated. Recently, experiments from our laboratory showed that in the goldfish pituitary, it is Smad3 that plays a major role in activin-stimulated FSHß expression at the promoter level, although both Smad2 and Smad3 are expressed in the pituitary (data not shown). It is interesting to note that Smad7 is also expressed in the oocytes. Although Smad7 is not directly involved in activin signaling, it functions as a negative regulator to block activin signal transduction from its receptors to Smad2 or Smad3 [13, 14], and its expression in the zebrafish oocytes provides a potential mechanism for an intracellular negative feedback on activin signaling. This, together with the abundant expression of follistatin in the oocytes, suggests that mechanisms exist at different levels of the follicle, both extracellularly and intracellularly, to keep the activin influence on the oocyte in check.

In summary, we demonstrate in the present study that activin subunits, activin-binding protein, activin receptors, and Smads are all expressed in the zebrafish ovarian follicle cells and that the entire activin signaling system is also expressed by the oocytes. These results provide strong support to our hypothesis that activin may promote zebrafish oocyte maturational competence by directly acting on the oocyte surface. Since activin subunits are predominantly expressed in the follicle cells rather than the oocytes, it is conceivable that activin molecules from the follicle cells regulate oocyte development in a paracrine fashion. Interestingly, in addition to expressing activin receptors and Smads for activin actions, the zebrafish oocytes also express high level of activin-binding protein follistatin. This suggests that although the oocytes are likely regulated by activin, they may actively participate in the regulation by releasing follistatin to modulate activin actions, and a closed-loop feedback involving activin and follistatin may exist in the zebrafish ovarian follicle between the follicle cells and the oocytes. More studies are needed to elaborate and substantiate this model (Fig. 10).



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  		FIG. 10. Hypothetical model for paracrine action of activin from the follicle layer on the oocyte in the zebrafish ovarian follicle


   FOOTNOTES
 
1 The work was substantially supported by grants (CUHK4176/99M, CUHK4150/01M and CUHK4258/02M) from the Research Grants Council of the Hong Kong Special Administrative Region to W.G. Back

2 Correspondence: Wei Ge, Department of Biology, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China. FAX: 852 2603 5646; weige{at}cuhk.edu.hk Back

Received: 27 June 2003.

First decision: 16 July 2003.

Accepted: 6 August 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Vale W, Rivier J, Vaughan J, McClintock R, Corrigan A, Woo W, Karr D, Spiess J. Purification and characterization of an FSH releasing protein from porcine ovarian follicular fluid. Nature 1986 321:776-779[CrossRef][Medline]
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