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BOR - Papers in Press, published online ahead of print July 13, 2005.
Biol Reprod 2005, 10.1095/biolreprod.105.041830
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BIOLOGY OF REPRODUCTION 73, 959–966 (2005)
DOI: 10.1095/biolreprod.105.041830
© 2005 by the Society for the Study of Reproduction, Inc.

Large-Scale Temporal Gene Expression Profiling During Gonadal Differentiation and Early Gametogenesis in Rainbow Trout1

Daniel Baron 3, Remi Houlgatte 4, Alexis Fostier 3, and Yann Guiguen 2, 3

Institut National de la Recherche Agronomique,3 INRA SCRIBE, IFR 140, Campus de Beaulieu, 35000 Rennes, France Institut National de la Santé et de la Recherche Médicale-INSERM,4 ERM 206, Parc Scientifique de Luminy, 13288 Marseille Cedex 09, France


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The overall understanding of the sex differentiation cascade in vertebrates is still growing slowly, probably because of the variety of vertebrate models used and the number of molecular players yet to be discovered. Finding conserved mechanisms among vertebrates should provide a better view of the key factors involved in this process. To this end, we used real-time reverse transcription-polymerase chain reaction to produce a temporal map of fluctuations in mRNA expression of 102 genes during sex differentiation and early gametogenesis in the rainbow trout (Oncorhynchus mykiss). We used these 102 temporal gene expression patterns as a basis for a hierarchical clustering analysis to find characteristic clusters of coexpressed genes. Analysis of some of these gene clusters suggested a conserved overall expression profile between the sex differentiation cascade in fish and mammals. Among these conserved molecular mechanisms, sox9, dmrt1, amh, nr5a1, nr0b1, igf1, and igf1ra are, for instance, characterized as early expressed genes involved in trout testicular differentiation as it is known or suggested in mammals. On the contrary, foxl2, fst, and lhr are characterized as early expressed genes during trout ovarian differentiation, as also found in mammals. Apart from this high conservation, our analysis suggests some potential new players, such as the fshb subunit gene, which is detected here for the first time, to our knowledge, in the female differentiating gonad of a vertebrate species and displays a specific overexpression that coincides in timing with the occurrence of first oocyte meioses, or the pax2 gene, which displays an early and testis-specific expression profile.

early development, gene regulation, ovary, testis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Sex differentiation, considered as the differentiation from a bipotential gonadal primordium toward a testis or an ovary, is a developmental process common to most species. A large research effort has focused on how some genes initiate and control cellular processes, such as cell division, migration, and differentiation, leading to testis or ovary formation in vertebrates [1].

In mammals, since the discovery of SRY [2], the Y-linked testis-determining gene, major involvement of several new genes in sex differentiation has been demonstrated. Analyses of these genes in humans showing gonadal dysgenesis or in mouse models, along with the use of in vitro cell culture assays, have revealed that the sex differentiation cascade results from a complex interplay between a large network of genes. Among these genes, special attention has been paid, for instance, to Sox9, Amh, Wt1, Nr5a1, Nr0b1, and Dmrt1 [3] in many vertebrate species, but nothing is really known about this overall gene network and its regulation.

In birds, reptiles, amphibians, and fish, extensive efforts have failed to find a functional orthologue to SRY, indicating an evolutionary plasticity in the sex determination switch (i.e., the initial switch that triggers the sex differentiation process). Despite this important difference with mammals, much information has been gathered on genes involved with the process of differentiation in lower vertebrates. Intriguingly, the sex specificity and the timing of expression for some of these genes during sex differentiation differ between species, but most of these genes are conserved and exhibit a characteristic expression during the period of sex differentiation of all these species.

In teleostean fish, sex-determining genetic systems are diversified, involving either a purely polygenic control, dominant sex-determining factors combined with autosomal controls, or a sex chromosome control, with heterogametic males in some species (XX/XY genetic system) or heterogametic females (ZZ/ZW genetic system) in others [4]. Environmental factors also may play a major role during sex determination in some of these species [5]. Except for the putative testis-determining factor, the dmy gene in the medaka (Oryzias latipes) [6], no sex-determining gene has been found in fish. Sex differentiation in fish can be controlled by in vivo treatments with sex steroids, as in reptiles and amphibians and, to some extent, in birds, and during the last few decades, most of the research involving fish has focused on the implication of these hormones in gonadal sex differentiation [7]. More recently, some new candidate genes have been shown to be involved in the fish sex differentiation cascade, and most of them are already known to be involved in the regulation pathways of vertebrate sex differentiation, including dmrt1 [8, 9], sox9 [10, 11], nr0b1 [12], amh [13, 14], and foxl2 [15].

The rainbow trout (Oncorhynchus mykiss) has a male heterogametic XY genetic system that is relatively insensitive to external factors. This makes it possible to produce males with new genotypes (XX and YY males) by hormonal sex inversion treatment and subsequent progeny testing [16]. When mated with normal females, these new male genotypes produce all-male (YY x XX) or all-female (XX x XX) populations [17]. These monosex populations have been widely used for studies of sex differentiation, because they give the opportunity to work on undifferentiated gonads for which the natural fate as testis or ovary is known a priori. However, apart from a few studies on a limited number of candidate genes, no investigations have been conducted on multigenes to understand the complexity of events that occur during gonadal sex differentiation in fish. Benefiting from some genome sequencing projects in trout [18–20], a large number of gene sequences either known or suspected to be involved in vertebrate sex differentiation are now directly available in public databases. In the present study, we took advantage of this resource and simultaneously analyzed the expression profiles of a large number of genes involved in vertebrate sex differentiation or early gametogenesis. These genes belong to the following main families: transcription factors, specific germ cell proteins, steroidogenic enzymes, hormones and growth factors, apoptosis regulators, and hormones and growth factor receptors. Expression profiles were obtained by real time reverse transcription-polymerase chain reaction (RT-PCR) that produces molecular signatures that are more accurate than those of other high-throughput methods, such as microarrays. These expression profiles were then analyzed using a standard hierarchical method generally employed for microarray data sets. Hierarchical clustering is widely used for cDNA array data analysis and is based on an overall processing of numerous gene profiles using algorithms that sort through all the data to find the pairs of genes that behave most similarly in all samples; it then progressively adds other genes to the initial pairs to form clusters of putatively coregulated genes. Although this method was developed initially to explore microarray results [21], we successfully applied it to our set of real-time RT-PCR expression values. The results obtained by real-time RT-PCR clearly clustered into biologically meaningful groups, revealing important molecular features of gonadal differentiation in fish and suggesting a high conservation of these molecular mechanisms with other vertebrates.


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

Research involving animal experimentation has been approved by the authors' institution (authorization no. 35-14). It conforms with principles for the use and care of laboratory animals and is in compliance with French and European regulations on animal welfare (European Convention for the Protection of Vertebrate Animals Used for Experimental and Other Scientific Purposes, ETS no. 123, 1 January 1991). All-male and all-female rainbow trout populations were obtained from the INRA experimental fish farm (Drennec, France) as previously described [17] using breeders from the Mirwart strain [22]. At 55-days postfertilization (i.e., the onset of free swimming), two batches of 1500 male and female fry were transferred to 0.3-m3 tanks with recirculating water conditions. They were held at 12°C under constant photoperiod (12L:12D) and were fed ad libitum with a commercial diet (dry pellet food; Biomar, Brande, Denmark). For each sex, 20–100 gonads, depending on the age of the fish, were sampled in duplicate at various stages of development and pooled. The following sampling dates were chosen: Day (D) 0 (onset of the free-swimming period after complete yolk resumption), D7, D12 (first oocyte meioses), D27 (first ovarian lamellar structures), D60 (first previtellogenic oocytes), D90, and D110. Samples were immediately frozen in liquid nitrogen and stored at –80°C until RNA extraction. Additional gonads were sampled at the same dates and fixed for histological analysis, which was performed as described previously [15].

Total RNA Extraction and RT

Total RNA was extracted using TRIzol reagent (Invitrogen, Cergy Pontoise, France) as described previously [23]. The total RNA concentration was determined with an Agilent 2100 Bioanalyzer and the RNA 6000 LabChip kit (Agilent Technologies, Stockport, UK) according to the manufacturer's instructions. For cDNA synthesis, 1 µg of RNA was denatured in the presence of random hexamers (0.5 µg) for 5 min at 70°C, then chilled on ice. Reverse transcription was performed at 37°C for 1 h using Moloney murine leukemia virus reverse transcriptase (Promega, Madison, WI) as described by the manufacturer.

Primer Design

Trout candidate gene homologues were sought in international public databanks using a reciprocal blast hit strategy [24]. Candidate genes were chosen according to a bibliographic analysis indicating their direct or indirect involvement in the sex differentiation cascade in vertebrates (Supplemental Table 1, available online at http://www.biolreprod.org). All primers were purchased from Eurogentec (Seraing, Belgium). These primers were manually designed and, whenever possible, respected the following restriction parameters: length, 21–23 base pairs (bp); no more than four successive identical nucleotides; guanin-cytosin content, 30–70%; a maximum of two guanin or cytosin among the five 3'-end bases; no primer dimmers; and a short amplicon size (70–150 bp). Secondary structures were sought using DNA mfold (http://bioinfo.math.rpi.edu/~mfold/DNA/form1.cgi) [25]. Whenever possible, each pair was chosen with at least one primer flanking an intron-exon boundary to prevent genomic amplification. This was not possible for some intronless genes, so we checked the amount of genomic contamination in all samples using an additional primer pair (up: GAGACAACCCTGGAGACCAA; down: CCAAGACCACCGCTTTAAGA) that was specifically designed to amplify genomic DNA from the tumor-suppressing gene tp53 (GenBank accession no. M75145). This genomic contamination was found to be very low or undetectable (cycle threshold, >37).

Real-Time RT-PCR

Real-time RT-PCR was carried out on an iCycler iQTM (Bio-Rad, Hercules, CA). Reactions were performed in 20 µl with 300 nM of each primer and 5 µl of a 1:50 dilution of the RT reaction and the SYBER-Green PCR Master Mix (Eurogentec) according to the manufacturer's instructions. After two incubation steps (50°C for 10 min, 95°C for 2 min), the thermal cycling protocol was as follows: 95°C for 10 min, followed by 40 cycles of PCR at 95°C for 30 sec and 60 or 65°C for 1 min. For each primer set (Supplemental Table 1, available online at http://www.biolreprod.org, the efficiency of the PCR reaction (linear equation: y = slope + intercept) was measured in triplicate on serial dilutions of the same cDNA sample (pool of reverse-transcribed RNA samples). Real-time PCR efficiencies (E)for each reaction were then calculated using the following equation: E = [10(1/slope)]–1. Melting-curve analysis also was performed for each gene to check the specificity and identity of the RT-PCR products. The relative amount of the target RNA, called the Starting Quantity (SQ), was then determined using the iCycler IQ software (Bio-Rad) by comparison with the corresponding standard curve for each sample run in duplicate. The SQ values were calculated as follows: SQ = , where Ct is the cycle threshold of the unknown sample. Each transcript level was then normalized by division with the expression values of the constitutive elongation factor 1{alpha} (ef1a), which was used as an internal standard.

Data Analysis

Gene expression profiles were used to classify genes and biological samples using a hierarchical clustering method with the Gene Cluster program [26]. The expression level of each gene was first log-transformed and then median-centered (relative to its median expression across all samples) so that relative variations rather than absolute values were used for interpretation. The clustering method employed was an average linkage with the Pearson correlation coefficient as a similarity metric. Results were displayed using the TreeView program [26]. To validate the most significant differences in gene expression profiles, we used the "Significance Analysis of Microarrays" (SAM) software [27]. Two comparisons were made. First, we performed a two-class unpaired analysis using male and female gene expression as classes, resulting in a list of significantly different gene profiles between sexes. Following an unsupervised clustering, we were able to further subdivide our samples into two other groups, described as early and late samples for both male and female profiles. We then ran a SAM analysis using a multiclass response to screen genes with significant expression profiles within these early or late, male or female groups. Finally, to discriminate only those genes that were differentially expressed in ovarian and testicular differentiating gonads, we ran a two-class unpaired analysis using only the two male and female early groups. For all SAM analyses, we used the lowest false-detection rate possible and the associated {Delta} value to determine the list of significant genes.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Histology

Males and females were surveyed over a 3-mo period starting from the time of first feeding (D0; complete yolk resumption and free swimming). Sampling dates were based on major events in gonadogenesis [17, 28]. Between D0 and D7, no difference could be observed in the germ line cells between male and female gonads (Fig. 1B). At that time, the gonadal ridge appears as a thin structure of coelomic epithelium in which gonia are proliferating. Around D12 to D27, the first occurrence of oocyte meiosis is detected in developing ovaries. Between D60 and D90, gametogenesis begins (Fig. 1B). Females develop a typical ovary; in its swollen anterior part, large previtellogenic oocytes are present in a lamellar structure. At the same time, males develop a testis characterized by an elongated, tiny structure in which cysts of dividing spermatogonia can be observed.



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  		FIG. 1. A) Classification of the biological samples using an unsupervised average linkage clustering of the gene expression profiles. B) Histological semithin sections of representative gonads. EC, epitheloid cell; G, gonia; I, interstitial tissue; L, lobule; M, mitosis; Me, meiosis; S, Sertoli cell; OL, ovarian lamellae; Oo, oocytes. Bar = 5 µm

Gene Expression Profiling

The complete data set is available via the National Center for Biotechnology Information (Bethesda, MD) Gene Expression Omnibus (GEO accession no. GSE2151), and the raw data set is included as online supplemental data (Supplemental Table 2, available online at http://www.biolreprod.org). For each sampling date, gene expression profiles were obtained on two independent pools of gonads (biological replicates), each containing 20 (D110) to 100 (D0) gonads, depending on their size. Two independent PCR measurements were performed for each RNA sample (technical replicates). To determine the consistency of our data, we calculated correlation coefficients (r) of both biological and technical replicates (Fig. 2, A and B). The average correlation values (r) of biological and technical replicates were on the same order of magnitude and ranged from 0.90 to 0.98 across the samples. Thus, our data are highly reproducible, presumably because the variation between biological replicate samples was minimized by the large number of individual gonads included in each RNA pool. On the contrary, no significant correlation could be detected between different biological samples. Thus, it can be assessed that measured variations reflect physiological differences (Fig. 2C).



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  		FIG. 2. Scatterplots of the 102 mRNA levels obtained (A) from two independent measurements of the same sample (female 1, D110), (B) from biological sample duplicates (female 1 vs. female 2, D110), and (C) from two different samples (female 1 vs. male 1, D110). Data are depicted with a logarithmic scale (log-log plots, base 10). Each black spot denotes a data point. The diagonal black line denotes the identity (log X = Log Y), and the gray lines indicate the twofold upper limit. Student t-tests (P = 0.05) were performed to compare the expression levels of genes between testis and ovary at D110 (C). Black spots with white bars correspond to genes significantly overexpressed in the ovary, and those with white dots correspond to genes significantly overexpressed in the testis. Particular genes are quoted on the graph as illustrations: pax2a (paired-box gene 2 a), igf1 (insulin-like growth factor 1), inha (inhibin {alpha} chain), fgf6 (fibroblast growth factor 6), fst (follistatin), fshb (ß subunit of follicle-stimulating hormone), bmp7 (bone morphogenetic protein 7), gdf9 (growth differentiation factor 9), wt1.2 (Wilms' tumor suppressor 1.2), and lama5 (laminin {alpha}5 chain)

Based on the efficiency and specificity of RT-PCR, 102 genes (see Supplemental Table 1, available online at http://www.biolreprod.org) were finally selected and analyzed. To assess the biological relevance of the genes assayed, we first performed an unsupervised hierarchical clustering (Fig. 1A) that clearly separated two large groups of samples, corresponding to male and female gonads. In each group, analysis of gene expression patterns discriminated two subgroups of samples: one that gathers the earliest samplings (D0, D7, D12, and D27), corresponding to differentiating gonads; and one (D60, D90, and D110) corresponding to the beginning of gametogenesis. Among all these groups, almost all the biological duplicates were clustered together, showing the efficiency of the method. Second, we ran a supervised clustering analysis with samples ordered on the basis of the development stage time (from D0 to D110) to highlight groups of correlated genes (Fig. 3). No invariant group was obtained, which is not surprising, because all genes were chosen on the basis of their differential expression during gonadal differentiation and/ or early gametogenesis. The SAM analysis (Fig. 3) showed that 74% of the genes displayed a statistically different expression profile in at least one of the four groups (early and late, male and female gonads). Clusters were identified according to sample classification using correlation thresholds of 0.6 and 0.8 for gene nodes. Analysis of the dendrogram reveals that genes are clustered into three main branches (threshold = 0.6). The first group corresponds to genes with the highest relative expression in the testis (node 1), the second group to genes with the highest relative expression in the ovary (node 2), and the third one to genes with similar expression profiles in the testis and the ovary (node 3). Using a threshold of 0.8, these groups can be further subdivided into five clusters named, respectively, C1.1 and C1.2 for node 1, C2.1 and C2.2 for node 2, and C3 for node 3. For each cluster, some representative expression profiles are shown in Figure 4.



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  		FIG. 3. Supervised average linkage clustering of the 102 gene expression profiles in rainbow trout male and female gonads. Each row represents a gene, and each column represents a sample. The 102 genes are referenced on the right according to the zebrafish nomenclature (see Supplemental Table 1, available online at http://www.biolreprod.org). The dendrogram on the left represents correlation distances between the profiles of the studied genes. The 28 samples at the top of the image are supervised according to sex (male and female) and the development stage time (D0 to D110). Each cell in the matrix corresponds to an expression level, with blue for overexpression, yellow for underexpression, and black for gene expression close to the median (see the color scale). The significant genes (SG) determined by the SAM analysis are represented on the right of the figure. A) Analysis of the four groups of samples discriminated by the unsupervised hierarchical clustering [75 SG, false-detection rate (FDR) = 0.209, Delta value ({Delta}) = 0.289, number of genes falsely called (GFC) = 0.157]. B) Analysis between male and female [41 SG, FDR = 0.574, {Delta} = 1.2, GFC = 0.235]. C) Analysis between early male and early female [35 SG, FDR = 1.568, {Delta} = 1.42, GFC = 0.549]. Blue asterisks indicate significant overexpression in male gonad, and red asterisks indicate significant overexpression in female gonad. Genes are grouped on the basis of similarity of their expression patterns across all samples, and particular clusters are quoted on the top right. *Significant difference



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  		FIG. 4. Expression profiles of some representative genes from the five main clusters (C1.1, C1.2, C2.1, C2.2, and C3) identified during male and female sex differentiation. Cluster C1.1: pax2a (paired-box gene 2 a) and igf1 (insulin-like growth factor 1); cluster C1.2: inha (inhibin {alpha} chain) and fgf6 (fibroblast growth factor 6); cluster C2.1: fst (follistatin) and fshb (ß subunit of follicle-stimulating hormone); cluster C2.2: bmp7 (bone morphogenetic protein 7) and gdf9 (growth differentiation factor 9); cluster C3: wt1.2 (Wilms' tumor suppressor 1. 2) and lama5 (laminin {alpha}5 chain). For each histogram, female samples are represented by black squares and male samples by open squares. Results are represented with an arbitrary scale as the ratios between the expression of the specific gene and ef1a. Each bar represents the mean of two different measurements (represented by black or white dots) from different biological samples (same male or female population, same sampling date)

Cluster C1.1 (Fig. 3) contains 22 genes, and among them, 19 exhibit a significantly higher relative expression in the testis compared to the ovary. Some of these genes display a roughly constant expression level from D0 to D110; such genes include amh, star, dmrt1, cyp11b2.1, cyp11b2.2, pax2a, sox9a.1, igf1ra, esr2b, igf1, and pdgfra (see pax2a and igf1 in Fig. 4). The other genes, namely igfbp2, nr0b1, inhba, nr5a1b, smad7, sox9a.2, gja1, and cyp11a1, show a marked underexpression in the ovary from D60 to D110 rather than an overexpression in the testis.

Cluster C1.2 (Fig. 3) contains 14 genes that are overexpressed in the late testicular group and display an almost median expression in the early and late ovarian groups. This cluster contains igf2, lhx9, inhbb, fshr, apoeb, hsd3b1, inha, stc1, cyp17a1, esr1, gata2, cldn11, fgf6, and tnfa (see inha and fgf6 in Fig. 4). Among these genes, fshr, apoeb, hsd3b1, inha, and stc1 are significantly underexpressed in the early testicular group compared to the early ovarian group. Finally, three genes (ara, arb, and bzrp) are clustered apart from C1.1 and C1.2.

Cluster C2.1 (Fig. 3) contains 12 genes that are overexpressed in the early ovarian group (see fst and fshb in Fig. 4). However, cyp19a1a, foxl2a and foxl2b, fst, lhr, ovol1, and hsd11b3 are highly expressed in all samples of the early ovarian group and decrease thereafter. On the contrary, bcl2lb, fshb, fancl, gcl, and lhb progressively increase from D0 to D27 and then decrease at D60 to reach levels roughly similar to those detected at D0. Cluster C2.2 (Fig. 3) is the largest, with 33 genes showing a late ovarian overexpression (from D60 to D110; see bmp7 and gdf9 in Fig. 4). Their ovarian expression is 10- to 20-fold higher at D110 than at D0. Some of these genes, namely vasa, cdh1, bmp7, vldr, bcl2la, and msh2, also display a similar pattern of overexpression in late testicular samples. The others are only overexpressed in the ovary. Among them, nup62, casp3, gdf9, sox24, lifr, aldob, tial1, and sox23 are overexpressed during the whole late ovarian period, and mapk8, vim, fgf2, ctnnb, birc5a, mapk3, hsd17b4, eif2s3, tgfbr2, bactin2, casp6, fasn, tra2a, hsp90b, magoh, and zp3a are only overexpressed slightly later (from D90 to D110). Finally, gapd, igf1rb, and ncor1 also belong to the node 2 (Fig. 3) but are clustered separately from C2.1 and C2.2.

Cluster C3 (Fig. 3) contains 21 genes showing a roughly similar pattern of expression, with no significant difference between the testicular and ovarian groups except for tgfb1 (see wt1.2 and lama5 in Fig. 4). These genes, namely tgfb1, cav1, solt, pdgfrb1, bad, creb1, wt1.1 and wt1.2, mtnr1ar, lama5, bmp4, ptgs2, akr1d1, tegt, ahr1, tfa, shbg, nr0b2, alox5, acvr1b, and timp2, show a higher expression in both early testicular and early ovarian groups and then decrease approximately twofold after D27. However, among these genes, bmp4, ptgs2, akr1d1, and tegt exhibit a marked underexpression in the late testicular group.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Emerging expressed sequence tag projects in rainbow trout [18–20] provide a new resource that enables us to study, both directly and simultaneously, numerous gene expression profiles. The present study emphasizes accurate temporal patterns based on the precision and reliability of real-time RT-PCR, but in comparison to the large number of gene expression profiles that could potentially be analyzed using chip technologies, we only produced 102 gene profiles. However, this number of genes is already too high to allow a "one by one" analysis. The overall expression patterns were then analyzed with a hierarchical clustering method and displayed in a color-coded matrix. This method has already been applied successfully to PCR data by Wen et al. [29] to highlight a group of coexpressed genes during development of the central nervous system. The sample classification we obtained using the clustering algorithm of Eisen et al. [26] is consistent with the genetic sex and with the time of sampling. This classification also clearly discriminates the male and female differentiating periods that are associated with specific clusters of genes for testicular and ovarian differentiation. This is noticeable because no sex difference is clearly observed in the germ cell lineage at this time.

Our results clearly show that male and female gonads, on the whole, share similar patterns of expression (21 genes) during the differentiating period (D0 to D27). Among these genes, some have been reported to be involved in cell proliferation, maintenance, and differentiation of the gonad, such as bmp4 and wt1. In mammals, Bmp4 is required for initiation [30] and proliferation [31] of the primordial germ cells in the embryonic gonad, and Wt1 (-kts isoform) is required for the survival and proliferation of somatic bipotential gonadal cells [32]. Other genes expressed during this period in the trout male or female gonads also have been reported to be involved in gonadal development of both sexes as structural macromolecules, such as the basement membrane Lama5 [33].

In addition to these genes sharing an overall similar pattern of expression in both sexes, we found numerous genes discriminating male and female gonads during the differentiating period (D0 to D27). In females, fst, ovol1, foxl2a, foxl2b, and lhr are over- and coexpressed with a well-known key factor of ovarian differentiation in fish, aromatase (cyp19a1a) [7]. Interestingly, in the chicken, lhr is overexpressed in the female gonad concomitantly with an aromatase overexpression [34], which may be related to an earlier production of estrogens in females, as we found in the present experiment. Expression of foxl2 in mammals [35, 36] starts before the occurrence of clear morphological features related to gonadal differentiation. This is consistent with our results, because the two duplicated trout foxl2 genes [15] are expressed at that time as well. Follistatin (fst) also exhibits a very early, specific ovarian expression in our experiment, and it appears to be a good candidate for promoting ovarian development. In mammals, Fst, which is necessary for the maintenance of ovarian germ cells, recently has been shown to inhibit formation of the XY-specific coelomic vessels in XX gonads [37], raising the possibility that the ovarian pathway operates by repressing the testicular pathway [38]. Another gene promoting ovarian development could be ovol1. In the fly, this gene is required for female germ line determination and differentiation [39]. Interestingly, Ovol1 also was found to be expressed in the sheep ovary before sex differentiation [40]. Additionally, homozygous Ovol1 –/– mice have been obtained. Unfortunately, the phenotypes were studied only in males, and no data are available on Ovol1 gene expression during ovarian differentiation in mammals [41].

We also found 20 genes that are specifically overexpressed during testicular differentiation. Among these genes, some have been widely shown to be involved in mammalian testis differentiation, such Sox9, Dmrt1, Nr5a1 (also called Sf1), Amh, and Nr0b1 (also called Dax1). Sox9 is up-regulated in the differentiating male gonad of most species and represents a key phenomena in all vertebrates regardless of the switch mechanism controlling sex determination [42]. Of great interest, Sox9 also has been shown to directly activate Amh expression with Nr5a1 [43]. The two genes are actually coexpressed with sox9a.1 and sox9a.2, which suggest that a similar regulation mechanism may exist in the differentiating trout testis. We also found an early testicular overexpression of nr0b1 (dax1) suggesting its implication in trout testicular differentiation. In mammals, its role is still controversial, because it was initially described as an ovary-determining candidate gene [44]. However, recent studies in the mouse have shown that it may, instead, be involved in the testicular differentiation pathway [45]. Additionally, dmrt1 was expressed specifically in the testis throughout our experiment. It has been shown previously to be involved in male differentiation in most species [46], including trout [9], but its precise role is still unknown. However, targeted deletion of mouse Dmrt1 revealed that this gene is essential not only for testis differentiation but also for testicular development [47], which is in agreement with our result, because its expression persisted until the end of the present experiment. We also found new genes that might be involved in trout testis differentiation. Particularly, igf1 and its receptor, igf1ra, were significantly overexpressed during testicular differentiation. Very interestingly, Igf1 has been shown to be essential for endocrine activity in the embryonic mouse testis [48], which is consistent with our results, because most steroidogenic enzymes and factors also are concomitantly overexpressed during this period in the differentiating trout testis. Even more recently, testis differentiation has been shown to require the insulin-receptor family function in mice, because XY males that are mutant for three receptors of the insulin family develop ovaries with a complete absence of Sox9 and Amh expression [49]. Finally, we found that a paired-box gene, pax2a, is specifically overexpressed in the differentiating trout testis. Interestingly, Pax2 is expressed in the mouse gonad [50] and is important to the formation and maintenance of the male reproductive tract in mammals [51], possibly interacting with another member of the family (Pax8) [51]. To our knowledge, however, no information is available on its putative role in the testis.

In trout, ovarian differentiation becomes morphologically evident when the first oocyte meioses are observed. Expression profiling revealed that some genes (bcl2lb, fancl, gcl, fshb, and lhb) are overexpressed specifically at this time during ovarian differentiation. This is the first time, to our knowledge, that fshb and lhb have been found to be expressed in the differentiating gonad. One possible role for these two hormones may be as antiapoptotic agents at the time of oocyte meioses. Such an effect of FSH has been suggested already [52]. That bcl2lb is expressed concomitantly could favor this hypothesis, because it has been shown to prevent gonocytes from apoptosis in the embryonic mouse gonad [53]. However, we failed to detect any significant cga (glycoprotein hormone {alpha} polypeptide) mRNA expression during that period in trout, and the functional significance of fshb and lhb expression would need to be further explored. However, such an expression of cga has been reported along with Fshb expression in mouse testis [54] and with Lhb in rat testis [55]. Even more recently, fshb, lhb, and cga have been shown to be synthesized de novo in oocytes of the adult gilthead seabream ovary [56]. Finally, the two genes Fancl and Gcl are essential for proper primordial germ cell migration and proliferation in the embryonic mouse gonad [57, 58]. Interestingly, gcl also has been shown recently to be specific to the ovary in the medaka [59].

During the early period of gametogenesis (D60 to D110), the ovarian and testicular expression profiles are clearly different, an observation that is in agreement with the morphological differences as detected by histology. The ovary consists of ovarian lamellae containing primordial follicles with a central oocyte (previtellogenic oocyte) surrounded by proliferating, flattened granulosa cells. Simultaneous with the development of these primordial follicles, we were able to characterize numerous genes specifically expressed in the ovary. Among these genes, sox23, sox24, and nup62 were specifically overexpressed, as shown previously in trout [60, 61]. The expression of gdf9, which also is high during this period, could be specifically localized in the previtellogenic oocytes, as shown in mammalian oocytes [62], and could activate star, which also is highly expressed during this period. On the other hand, the flattened granulosa cells that proliferate at this time to finally surround the oocytes could express genes, such as ctnnb and bmp7, that are overexpressed in the trout ovary at this time as well and that have been shown to be expressed in granulosa cells of other species [63, 64]. That expression of ctnnb and bmp7 strongly increases simultaneously with the granulosa cell proliferation between D90 and D110 is in agreement with this hypothesis.

At the same time, males develop a testis with spermatogonia surrounded by Sertoli cells and organized into cysts. Some of the genes that we found to be specifically overexpressed in the testis at this time already have been shown to be highly expressed in the trout testis, such as fgf6 [65] and inha [66]. Others, such as cldn11, are well known as markers of the differentiated testis in mammals. In the mouse, Cldn11 is specifically expressed in Sertoli cells [67]. The situation might be similar in trout as well, because its overexpression is concomitant with Sertoli cell differentiation.

Despite the anatomical differences between a testis and an ovary during early gametogenesis, some common expression profiles can be observed, such as vasa and bmp7 genes. The vasa gene has been shown previously to be closely related to germ cells in trout [68]. Interestingly, Bmp7 also has been shown to be strongly expressed in germ cells in the mouse testis [69] and could play such a role in trout as well.

In conclusion, this original strategy of high-throughput, real-time RT-PCR gene profiling combined with an overall hierarchical clustering analysis produces the first comprehensive view of what is going on during rainbow trout gonadal sex differentiation. This analysis revealed that in terms of gene expression, sex differentiation in trout shares strong similarities with what has already been reported in other vertebrate species, including mammals. Apart from this overall conservation of gene expression, the present work also brings putative new players into the cascade of sex differentiation and early gametogenesis in trout that could themselves also be conserved in other vertebrate species. New investigations are currently underway to further decipher gene regulation during trout gonadal sex differentiation using high-throughput chip technologies.


   ACKNOWLEDGMENTS
 
The authors acknowledge Dr. Corine Cotinot for her advice in listing candidate genes. They also are grateful to the experimental facility staff of the INRA-SCRIBE laboratory for taking care of fish rearing and thank Robert Britton for correction of the manuscript.


   FOOTNOTES
 
1 Supported by funds from the "Institut National de la Recherche Agronomique" (INRA), from PNETOX, and from CIPA-OFIMER-INRA (IFOP) grants. D.B. received a fellowship from the "Ministère de la Recherche et de l'Enseignement Supérieur." Back

2 Correspondence: Guiguen Yann, INRA SCRIBE, Campus de Beaulieu, 35000 Rennes, France. FAX: 33 2 23 48 50 20; Yann.Guiguen{at}rennes.inra.fr Back

Received: 10 March 2005.

First decision: 14 April 2005.

Accepted: 5 July 2005.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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