Abstract
Changes in zebrafish testicular gene expression induced by follicle-stimulating hormone (Fsh) or anti-Mullerian hormone (Amh) suggested that Amh inhibition and Fsh stimulation of spermatogenesis involved up and downregulation, respectively, of prostaglandin (PG) signaling. We found that Sertoli cells contacting type A undifferentiated (Aund) and differentiating (Adiff) spermatogonia expressed a key enzyme of PG production (Ptgs2); previous work showed that Sertoli cells contacting Adiff and B spermatogonia and spermatocytes showed ptges3b expression, an enzyme catalyzing PGE2 production. In primary testis tissue cultures, PGE2, but not PGD2 or PGF2α, reduced the mitotic activity of Adiff and their development into B spermatogonia. Vice versa, inhibiting PG production increased the mitotic activity of Adiff and B spermatogonia. Studies with pharmacological PG receptor antagonists suggest that an Ep4 receptor mediates the inhibitory effects on the development of spermatogonia, and cell-sorting experiments indicated this receptor is expressed mainly by testicular somatic cells. Combined inhibition of PG and steroid production moreover reduced the mitotic activity of Aund spermatogonia and led to their partial depletion, suggesting that androgens (and/or other testicular steroids), supported by PGE2, otherwise prevent depletion of Aund. Androgens also decreased testicular PGE2 production, increased the transcript levels of the enzyme-catabolizing PGs and decreased PGE2 receptor ptger4b transcript levels. Also Fsh potentially reduced, independent of androgens, PGE2 production by decreasing ptges3b transcript levels. Taken together, our results indicate that PGE2, via Ep4 receptors, favors self-renewal in conjunction with androgens and, independent of Fsh and androgens, inhibits differentiating divisions of spermatogonia.
Introduction
Prostaglandins (PGs) are low-molecular-weight (~350 Da), polyunsaturated lipophilic signaling molecules. They are derived from arachidonic acid, which can be liberated from membrane phospholipids by phospholipase A2 (PLA2) enzymes. For PG biosynthesis, arachidonic acid is metabolized by cyclooxygenase enzymes 1 and 2 (COX 1 and 2, a.k.a. PTGS (prostaglandin synthase) 1 and 2) to PGH2, which is converted further by specific synthases, for example, PGD synthase (PTGDS) or PGE synthase (PTGES), to produce the main bioactive PGs (i.e. PGE2, PGD2, PGF2α, PGI2 (Simmons et al. 2004, Ricciotti & FitzGerald 2011)). PGs develop biological activity by interacting with one of the prostanoid receptors, a group of G-protein-coupled membrane receptors showing five subtypes with in total nine receptors (D1 and 2, E1-4, FP, IP and TP (Breyer et al. 2001, Bos et al. 2004)).
PLA2, COX1/2 and PG-specific synthases as well as prostanoid receptors are expressed by different tissues and cell types. It is therefore not surprising that PGs influence in a paracrine and/or autocrine manner several physiological systems including the CNS, cardiovascular, gastrointestinal, excretory, respiratory, immune, reproductive and endocrine systems (Hata & Breyer 2004). The half-life time of PGs measures in minutes in mammals (Shrestha et al. 2012). Despite this short half-life time, there is a specific dehydrogenase inactivating PGs (HPGD). Its physiological relevance is exemplified by the several phenotypes that have been associated with mutations in this gene in humans (https://www.genecards.org/cgi-bin/carddisp.pl?gene=HPGD), highlighting the wide range of processes modulated by PGs.
Inhibitors of COX1/2 activities block the biosynthesis of all PGs and are known as non-steroidal anti-inflammatory drugs (NSAIDs) (Hata & Breyer 2004). Early in vivo experiments indicated that treatment with NSAIDs or different PGs modulated spermatogenesis in mammals (e.g. mouse (Abbatiello et al. 1975, 1976) and dog (Moskovitz et al. 1987)). Later studies in rat suggested that part of these effects reflect PG-mediated modulation of Leydig cell steroidogenesis, considering that PDG2-promoted while PGF2α suppressed androgen production (Frungieri et al. 2015). In rat, FSH stimulated Sertoli cell PG production (Jannini et al. 1994). In fetal mice, Sertoli cell-derived PGD2 keeps spermatogonia in an undifferentiated state by autocrine stimulation of CYP26B1 activity that inactivates retinoic acid (Rossitto et al. 2015). Moreover, PGD2 triggered changes in gene expression in spermatogonia, reduced their cell cycling activity and upregulated Nanos2, in turn preventing Stra8 expression, which contributed to keep spermatogonia in an undifferentiated state in fetal mice (Moniot et al. 2014). PG effects on stem cells have also been described in certain malignancies, such as bladder cancer (Kurtova et al. 2015), where blocking PGE2 production prevented cancer stem cell self-renewal proliferation between chemotherapy cycles. A role for PGE2 in promoting stem cell production was also reported from non-malignant tissue, such as the hematopoietic stem cells in zebrafish (Choudhuri et al. 2017).
A number of reproductive processes in adult teleost fish have been reported to be influenced by PGs, such as sex steroid production (Wade & Van der Kraak 1993) or ovulation and reproductive behavior (Sorensen & Goetz 1993, Takahashi et al. 2018). With respect to an early life-stage during zebrafish sex differentiation, it is interesting to note that PGD2 resulted in male-biased, PGE2 in female-biased sex ratios (Pradhan & Olsson 2014). However, we are not aware of studies on the role of PGs in adult spermatogenesis in fish. In our studies on the endocrine and paracrine regulation of adult spermatogenesis in zebrafish, we have focused on the mitotic phase of spermatogenesis. This included gene expression analyses of zebrafish testis tissue exposed to recombinant hormones and growth factors (Crespo et al. 2016, Morais et al. 2017). We found that Fsh downregulated testicular ptgs2a, encoding Cox2a (Crespo et al. 2016), while this transcript was upregulated by Amh (Morais et al. 2017); Amh also upregulated the PGE receptor 4b transcript ptger4b, while Fsh decreased the expression of the PGE synthase ptges3b (Crespo et al. 2016). These data set suggest that Fsh (promoting spermatogenesis) downregulated PGE2 signaling, while Amh (inhibiting spermatogenesis) promoted PGE2 signaling. Our first experiments to test this assumption indeed showed that in the presence of Fsh, PGE2 decreased the proliferation activity of type A differentiating (Adiff) and type B spermatogonia (Morais et al. 2017). Here, we report our follow-up studies on the role of PGs in zebrafish spermatogenesis. Our data suggest that PGE2 production and signaling is part of the testicular regulatory network implementing the endocrine control of spermatogenesis and functions to prevent depletion of Aund spermatogonia while dampening both basal and Fsh- or 11-ketotestosterone (11-KT)-stimulated differentiation of spermatogonia.
Materials and methods
Animals
Wild-type and transgenic Tg(vasa:EGFP) adult male zebrafish (Danio rerio) between 4 and 12 months of age were used in the present study. Animal housing and experimentation were consistent with the Dutch national regulations. The Life Science Faculties Committee for Animal Care and Use in Utrecht (The Netherlands) approved the experimental protocols, which were carried out under permission NVWA 10800.
Testis tissue culture
A previously established testis tissue culture system (Leal et al. 2009b ) was used to investigate the involvement of prostaglandins and related compounds in zebrafish spermatogenesis including PGE2, PGD2, PGF2α, indomethacin (INDO), GW627368X (EP4 receptor antagonist) and PF04418948 (EP2 receptor antagonist; all purchased from Cayman Chemical) and used at a final concentration of 5 μM (except for INDO and PF04418948, which were used at 3 μM). Additional testis tissue culture experiments were carried out to investigate the interaction of PGE2 with the androgen 11-ketotestosterone (200 nM (Skaar et al. 2011); Sigma-Aldrich) or recombinant zebrafish Fsh (100 ng/mL (Nobrega et al. 2015)). To study PGE2 and/or Fsh effects in the absence of steroid hormones, incubations were carried out in the presence of trilostane (Tril, 25 μg/mL (Garcia-Lopez et al. 2010); Sigma-Aldrich), which prevents the production of biologically active steroids.
Morphometric analyses
To evaluate the proportion of area occupied by different germ cell types, testis tissue was fixed in 4% glutaraldehyde (4°C, overnight), dehydrated, embedded in Technovit 7100 (Kulzer), and 4 µm thick sections were stained with toluidine blue. Ten to fifteen randomly chosen, non-overlapping fields were photographed (Olympus AX70 microscope; ×400 magnification), and the images were analyzed quantitatively using a plugin of the ImageJ software, as previously described (Assis et al. 2016). The germ cell types analyzed (type A undifferentiated, Aund; A differentiating, Adiff; and B spermatogonia) were identified according to previously published morphological criteria (Leal et al. 2009a ).
Immunohistochemistry (IHC)
After fixation of testis tissue from adult zebrafish in 4% paraformaldehyde and embedding in paraffin, 4 µm thick sections were prepared for fluorescent IHC. Briefly, slides were heated for 10 min in sodium citrate buffer pH 6.0 for antigen retrieval and subsequently cooled to room temperature for 30 min. After washing in PBS and blocking with 5% BSA (in PBS), slides were incubated with a primary goat anti-(human)COX2 (a.k.a. PTGS2) polyclonal antibody (Cayman Chemical; previously used in zebrafish (Feng et al. 2012)), diluted 1:100 in 1% BSA/PBS. Then, slides were incubated with the secondary antibody (chicken anti-goat IgG Alexa Fluor 488; Life Technologies) for 90 min. Propidium iodide (Sigma-Aldrich) was used as nuclear counterstain and slides were mounted with Vectashield H-1000 (Vector Laboratories). Testis sections were analyzed using a confocal laser scanning microscope (Zeiss LSM 700).
The proliferation activity of type A and B spermatogonia was investigated by quantifying the cell fraction incorporating the S-phase marker bromodeoxyuridine (BrdU; 50 µg/mL, Sigma-Aldrich), which was added to the medium for the last 6 h of tissue culture. After incubation, testis tissue was fixed at room temperature for 1 h in freshly prepared methacarn (60% (v/v) methanol, 30% chloroform and 10% acetic acid glacial; Merck Millipore) and processed for subsequent analysis. Testis tissue was dehydrated, embedded in Technovit 7100 (Heraeus Kulzer), sectioned at a thickness of 4 μm. BrdU immunodetection was carried out as previously described (Leal et al. 2009a ). To quantify spermatogonial proliferation, the mitotic index was determined by examining at least 100 germ cells/cysts, differentiating between BrdU-labeled and unlabeled cells.
PGE2 and 11-KT measurements by ELISA
The levels of PGE2 produced by testis explants were analyzed directly in the incubation media using a commercial immunoassay (Prostaglandin E2 ELISA Kit-Monoclonal; Cayman Chemical). The levels of 11-KT in the incubation media were analyzed by ELISA, as previously described (Cuisset et al. 1994). Acetylcholine esterase-labeled tracers and microplates pre-coated with monoclonal mouse anti-rabbit IgG were supplied by Cayman Chemicals. Anti-11-KT was a kind gift from David E Kime (Sheffield University, UK). All samples were analyzed in duplicate.
Testicular gene expression
After tissue culture, testis tissue was frozen in liquid N and stored at −80°C until RNA isolation. Total RNA extraction, cDNA synthesis and real-time quantitative PCR (qPCR) were carried out as described previously (Nobrega et al. 2015). Data were normalized to eef1a1l1 (eukaryotic translation elongation factor 1 alpha 1, like 1) due to its stable expression in all sample groups analyzed. All qPCRs were performed in 20 µL reactions and quantification cycle (Cq) values were obtained in a Step One Plus Real-Time PCR system (Applied Biosystems) using default settings. Relative mRNA levels were calculated as reported previously (Bogerd et al. 2001). Primers used in the qPCR analyses are listed in Table 1.
Primers used for gene expression studies by qPCR analysis.
| Target gene | Gene description | Primer sequence (5′→3′) |
|---|---|---|
| amh | anti-Müllerian hormone | CTCTGACCTTGATGAGCCTCATTT GGATGTCCCTTAAGAACTTTTGCA |
| ar | androgen receptor | ACGTGCCTGGCGTGAAAA CAAACCTGCCATCCGTGAAC |
| cyp17a1 | cytochrome P450, family 17, subfamily A, polypeptide 1 | GGGAGGCCACGGACTGTTA CCATGTGGAACTGTAGTCAGCAA |
| dazl | deleted in azoospermia-like | AGTGCAGACTTTGCTAACCCTTATGTA GTCCACTGCTCCAAGTTGCTCT |
| eef1a1l1 | eukaryotic translation elongation factor 1 alpha 1, like 1 | GCCGTCCCACCGACAAG CCACACGACCCACAGGTACAG |
| foxa2 | forkhead box A2 | GTCAAAATGGAGGGACACGAAC CATGTTGCTGACCGAGGTGTAA |
| gsdf | gonadal somatic cell derived factor | CATCTGCGGGAGTCATTGAAA CAGAGTCCTCCGGCAAGCT |
| hpgd | 15-hydroxyprostaglandin dehydrogenase | GAGTAAAGAGTACGGAAAGCAAGGA GGTGAGGAGAATGGAGAAAAGCT |
| igf3 | insulin-like growth factor 3 | TGTGCGGAGACAGAGGCTTT CGCCGCACTTTCTTGGATT |
| insl3 | insulin-like 3 (Leydig cell) | TCGCATCGTGTGGGAGTTT TGCACAACGAGGTCTCTATCCA |
| piwil1 | piwi-like protein 1 | GATACCGCTGCTGGAAAAAGG TGGTTCTCCAAGTGTGTCTTGC |
| ptger2a | prostaglandin E receptor 2a (subtype EP2) | CTGTGGTTCAAACGGCGTATTT ACACACCGCATGAGTCTTGCT |
| ptger4a | prostaglandin E receptor 4 (subtype EP4) a | GCGGAGATCCAGATGGTCAT TGGGTTTTTATCCAGACGCTTCT |
| ptger4b | prostaglandin E receptor 4 (subtype EP4) b | GTGCTCATCTGCTCCACTCCTT GCAGAGTTAAACAGCTGGTTCACA |
| ptges | prostaglandin E synthase | GCCAAGTGAGACTTCGGAAAAA AACTGCACACCTCCGTGTCTCT |
| ptgesl | prostaglandin E synthase 2-like | GCAATTCATGGGAGGTGATGA TCCATAACCCTCAGAACTCCAAAC |
| ptges3b | prostaglandin E synthase 3b (cytosolic) | GACAGCAAAGACGTGAAAGTAAATTTT CGGCTCCACTGAGACAGCTAA |
| ptgs1 | prostaglandin-endoperoxide synthase 1 | ATTCAATCTGAAACCCTACACATCCT CGTATAGTTCCTCCAGCTCTTTAGACA |
| ptgs2a | prostaglandin-endoperoxide synthase 2a | ACGCTGGAGGTTCAACACAAA CACCTGGACGTCCTTCACAAG |
| rpl13a | ribosomal protein L13a | GAGCCCCCAGCAGAATCTTC AGCCTGACCCCTCTTGGTTTT |
| star | steroidogenic acute regulatory protein | CCTGGAATGCCTGAGCAGAA ATCTGCACTTGGTCGCATGAC |
| ubc | ubiquitin C | CCATACACCGCACTCTTACAGAAA CCAGTCAGCGTCTTCACAAAGAT |
| wnt5a | wingless-type MMTV integration site family, member 5a | TGGAGATCGTGGACGCAAA CACTTCAGGAATCAGCAGAGGATT |
Fw, forward; Rv, reverse.
To investigate the cellular expression of Ep2a and Ep4b receptors in the testis, the transcript levels of ptger2a and ptger4b, respectively, were retrieved from an RNA sequencing (RNAseq) dataset available in our group and submitted to the NCBI GEO database (GSE116611). This set contains expression data from control testes, from germ cell-depleted testes following treatment with the cytotoxic agent busulfan, and from testes with spermatogenesis recovering from the busulfan treatment. A complete description of this data set, involving the three experimental groups, will be given elsewhere. The data – also used to examine testicular PG-related gene expression in the untreated control group shown in Table 2 – were generated, assembled and analyzed as previously described (Morais et al. 2017). Briefly, RNAseq sequencing libraries (five biological replicates per treatment) were prepared from 2 µg total RNA using the Illumina TruSeq RNA Sample Prep Kit v2 and sequenced on an Illumina HiSeq2500 sequencer (Illumina, Inc.) as 1 × 50 nucleotide reads. Data analysis was performed using the R/Bioconductor package DESeq.
Prostaglandin-related gene expression in the adult zebrafish testis.
| Gene ID | Gene description | Gene symbol | Read number |
|---|---|---|---|
| ENSDARG00000052148 | prostaglandin-endoperoxide synthase 1 | ptgs1a | 87.4 |
| ENSDARG00000004539 | prostaglandin-endoperoxide synthase 2a | ptgs2aa | 62.3 |
| ENSDARG00000010276 | prostaglandin-endoperoxide synthase 2b | ptgs2b | 32.5 |
| ENSDARG00000069439 | prostaglandin D2 synthase a | ptgdsa | 9.9 |
| ENSDARG00000027088 | prostaglandin D2 synthase b, tandem duplicate 1 | ptgdsb.1 | 231.2 |
| ENSDARG00000071626 | prostaglandin D2 synthase b, tandem duplicate 2 | ptgdsb.2 | 88.5 |
| ENSDARG00000078605 | prostaglandin E receptor 1a (subtype EP1) | ptger1a | 12.9 |
| ENSDARG00000078602 | prostaglandin E receptor 1c (subtype EP1) | ptger1c | 2.4 |
| ENSDARG00000011434 | prostaglandin E receptor 2a (subtype EP2) | ptger2aa | 28.9 |
| ENSDARG00000037033 | prostaglandin E receptor 2b (subtype EP2) | ptger2b | 8.9 |
| ENSDARG00000055781 | prostaglandin E receptor 3 (subtype EP3) | ptger3 | 12.8 |
| ENSDARG00000059236 | prostaglandin E receptor 4 (subtype EP4) a | ptger4aa | 84.9 |
| ENSDARG00000035415 | prostaglandin E receptor 4 (subtype EP4) b | ptger4ba | 7.6 |
| ENSDARG00000079907 | prostaglandin E receptor 4 (subtype EP4) c | ptger4c | 12.1 |
| ENSDARG00000020136 | prostaglandin E synthase | ptgesa | 24.0 |
| ENSDARG00000068415 | prostaglandin E synthase 2-like | ptgesla | 756.8 |
| ENSDARG00000037284 | prostaglandin E synthase 3a (cytosolic) | ptges3a | 1872.1 |
| ENSDARG00000089626 | prostaglandin E synthase 3b (cytosolic) | ptges3ba | 16.9 |
| ENSDARG00000074016 | prostaglandin F receptor (FP) | ptgfr | 2.6 |
| ENSDARG00000078172 | prostaglandin F2 receptor inhibitor | CU984600.1 | 48.1 |
| ENSDARG00000060094 | prostaglandin I2 (prostacyclin) synthase | ptgis | 51.1 |
aIndicates genes for which a qPCR system was developed (as shown in Table 1).
To examine if ptger2a and ptger4b are expressed in somatic or in germ cells, we also used testes from transgenic Tg(vasa:EGFP) zebrafish expressing enhanced green fluorescent protein (EGFP) under the control of the germ cell-specific vasa promoter (Krovel & Olsen 2002). Testicular cell suspensions were prepared as described previously (Hinfray et al. 2013) and immediately submitted to fluorescence-activated cell sorting (FACS) using an in Flux cell sorter (BD Bioscience). EGFP-positive and -negative cells were collected, centrifuged in PBS at 100 g for 10 min and the pellet stored at −80°C until use for gene expression analysis by qPCR.
Statistical analysis
GraphPad Prism 5.0 package (GraphPad Software, Inc.) was used for statistical analysis. Significant differences between groups were identified using Student’s t test (paired or unpaired, as appropriate) or one-way ANOVA followed by Tukey’s test for multiple group comparisons (*P < 0.05; **P < 0.01; ***P < 0.001). Data are represented as mean ± s.e.m.
Results
Prostaglandin-related gene and protein expression in the zebrafish testis
Analysis of RNAseq data obtained using testis samples of untreated adult zebrafish (data set GSE116611) revealed expression of numerous PG-related genes, such as key enzymes involved in the production of all PG series (ptgs1, ptgs2a, ptgs2b; Table 2). The majority of transcripts retrieved were associated to PGE2 signaling (including several receptors and synthases), but their expression levels often were low (<15 normalized reads for 6 out of 12 transcripts identified; Table 2), compared to the 31,855, 7223 and 14,203 reads sequenced for three selected housekeeping genes eef1a1l1, rpl13a and ubc, respectively (data set GSE116611). Additional receptors and synthases for other PG types (i.e. PGD2, PGF2α and PGI2; Table 2) were also detected.
In order to investigate which cell type(s) are involved in testicular PG production, an fluorescent immunodetection approach for Ptgs2 was used. Ptgs2 labeling was observed in Sertoli cells, particularly in the nuclei of those Sertoli cells supporting type A (both Aund and Adiff) spermatogonia (Fig. 1A, B, C, D, E and F). Pre-adsorbing the antibody with the peptide sequence used for its generation eliminated the staining (Fig. 1G, H and I), showing that unspecific staining did not contribute to the labeling pattern observed.
PGs are produced by Sertoli cells in adult zebrafish testes. (A, B, C, D, E and F) Detection of Ptgs2 protein by fluorescent immunohistochemistry. Green staining indicates Ptgs2-positive Sertoli cells and red staining indicates DNA (propidium iodide counterstain). (G, H and I) Ptgs2 antibody pre-adsorbed with blocking peptide as negative control. Arrowheads indicate nuclear Sertoli cell (SC) staining, and representative type A undifferentiated (Aund) and differentiating (Adiff) spermatogonia are encircled with a white dashed line. Scale bars in C, F and I represent 10 µm. Ptgs2-positive nuclei were restricted to Sertoli cells contacting type Aund and Adiff spermatogonia. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0309.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
Blocking testicular PG production increased spermatogonial proliferation and differentiation
To investigate the possible involvement of PGs in regulating zebrafish spermatogenesis, we incubated testicular explants in the absence or presence of a potent PG production inhibitor (indomethacin; INDO). First, we examined if INDO treatment efficiently inhibited testicular PGE2 production. After both 1 and 4 days of incubation, PGE2 levels in the culture medium were significantly lower in the presence of INDO (Fig. 2A). Blocking testicular PG production elevated the proliferation activity of differentiating (type Adiff and B) spermatogonia (Fig. 2B), associated with an increase in the proportion of area occupied by type B spermatogonia (Fig. 2C). The expression of steroidogenesis-related genes (star and cyp17a1) was significantly reduced by INDO, while the other transcripts analyzed (germ cell markers and growth factors) were not affected by the treatment (Fig. 2D). Moreover, transcript levels of several PG-related genes (ptgs1, ptgs2a, ptges, ptgesl, ptges3b, ptger2a, ptger4a, ptger4b) did not change in response to INDO (data not shown). Hence, blocking PG production facilitated the proliferation of differentiating spermatogonia.
Inhibition of PG production by INDO promotes the differentiation of zebrafish spermatogonia. (A) Quantification of PGE2 production by testis tissue cultured for 1 or 4 days in basal medium or in medium containing 3 µM INDO (i.e. indomethacin, a nonsteroidal anti-inflammatory drug). Data are expressed as mean ± s.e.m. (n = 4–6) and asterisks indicate significant differences between groups (*P < 0.05, ***P < 0.001, paired Student’s t test). Additional testicular explants were cultured for 4 days in the absence or presence of 3 µM INDO and then used to quantify BrdU labeling indices (B), the areas occupied by type A and B spermatogonia (C), or candidate gene expression (D). Data are expressed as mean ± s.e.m. (n = 6–8) and asterisks indicate significant differences between groups (*P < 0.05, **P < 0.01, paired Student’s t test). In D, results are shown relative to the basal control condition, which is set at 1 (dashed line). Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
In view of the reduced levels of steroidogenesis-related transcripts following INDO treatment, we investigated possible interactions between PG and steroid signaling. First, we examined if the response to INDO is modulated when blocking the production of biologically active steroids with trilostane (Tril). Similar to the previous experiments (Fig. 2B and C), the differentiating spermatogonia showed higher proliferation rates and occupied increased proportions of the testicular area (Fig. 3A and B, respectively). In addition, Aund spermatogonia showed a lower proliferation activity and occupied a smaller proportion of the section surface. Also, lower transcripts levels of foxa2 (a potential marker for Aund spermatogonia (Safian et al. 2017)) and ar were observed under these conditions (i.e. INDO plus Tril; Fig. 3C), while the level of growth factor transcripts remained unaltered. Interestingly, the expression level of genes with products related to steroidogenesis remained unchanged, so that the PGE2 effect on star and cyp17a1 transcript levels observed earlier (Fig. 2C) may depend on the presence of steroids. In summary, the proliferation of differentiating spermatogonia was still favored when blocking PG and steroid production, while type Aund spermatogonia became partially depleted. However, type Aund spermatogonia remained unaffected when blocking only PG production.
INDO-induced differentiation, but not self-renewal, of spermatogonia is unaffected when blocking steroid production. Determination of BrdU labeling indices (A) and frequencies (B) of type A and B spermatogonia, and (C) quantification of candidate gene expression. Testicular explants were cultured for 4 days in the absence or presence of 3 µM INDO and collected for further analysis. The medium contained trilostane (Tril, 25 µg/mL) to block the production of biologically active steroids. Data are expressed as mean ± s.e.m. (n = 6–8). Asterisks indicate significant differences between groups (*P < 0.05, **P < 0.01, ***P < 0.001, paired Student’s t test). In C, results are shown relative to the basal control condition, which is set at 1 (dashed line). Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
PGE2 inhibits androgen- or Fsh-stimulated spermatogonial differentiation but promotes their self-renewal when combined with androgen
In order to study the effects of PGs on the proliferation of spermatogonia in a more direct manner, we added PGE2 to the culture medium, asking if PGE2 inhibits androgen-driven spermatogonia development. We found that PGE2 reduced the proliferation activity and area occupied by type B spermatogonia (Fig. 4A and B). Type Aund spermatogonia, on the other hand, showed an increased proliferation activity and accumulation, suggesting that these cells preferentially underwent self-renewal divisions in the presence of 11-KT and PGE2. Under these conditions, transcript levels of markers for differentiating spermatogonia (piwil1 and – not significantly – dazl) were downregulated, and considering growth factor expression, wnt5a levels increased (Fig. 4C). PGE2 treatment reduced two genes with products related to steroidogenesis (star and ar; Fig. 4C), but testicular androgen (11-KT) release did not change significantly (left panel in Fig. 4D). Conversely, 11-KT reduced PGE2 release (right panel in Fig. 4D), increased transcripts levels of the PG metabolizing enzyme hpgd (Fig. 4E), and modulated PG receptor (but not PG synthesis) gene expression (Fig. 4E).
In the presence of the androgen 11-KT, PGE2 increases the proliferation of type Aund spermatogonia while reducing their further differentiation. Testis tissue was cultured for 4 days in the absence or presence of 5 µM PGE2, in medium containing 200 nM 11-KT, and then used for quantification of BrdU labeling indices (A), areas occupied by type A and B spermatogonia (B), or candidate gene expression (C). Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia. (D) Quantification of 11-KT and PGE2 in testis tissue medium after culture for 1 day under different conditions. Different letters indicate significant differences between groups. (E) Modulation of PG-related gene expression by 11-KT. Results are expressed as mean ± s.e.m. (n = 6–17) and asterisks indicate significant differences between groups (*P < 0.05, **P < 0.01, paired Student’s t test). In C and E, results are shown relative to the basal control condition, which is set at 1 (dashed line).
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
Next, we examined the effect of PGE2 on Fsh-stimulated spermatogonia development. To focus on the steroid-independent effects of Fsh, trilostane (Tril) was included in the incubation medium. Similar to the findings in the presence of androgen, addition of PGE2 reduced Fsh-stimulated proliferation activity and relative numbers of differentiating spermatogonia (Fig. 5A and B). The effect of other PG types (PGD2 and PGF2α) on the proliferation activity were also evaluated in the presence of Fsh, but no significant changes were found for any of the three types of spermatogonia analyzed (data not shown).
PGE2 inhibits Fsh-stimulated differentiation of spermatogonia. Testis explants were cultured for 4 days with Fsh (100 ng/mL) in the absence or presence of 5 µM PGE2, while inhibiting steroid production with trilostane (Tril, 25 μg/mL). After 4 days, testis tissue was used for quantification of BrdU labeling indices (A), the areas occupied by type A and B spermatogonia (B), or of candidate gene expression (C). Quantification of 11-KT and PGE2 (D) in testis tissue medium after culture for 1 day under different conditions. Results are expressed as mean ± s.e.m. (n = 4–9) and asterisks indicate significant differences between groups (*P < 0.05, **P < 0.01, paired Student’s t test (A, B, C, and left panel in D) or one-way ANOVA followed by Tukey’s multiple comparison test (right panel in D). In (C), results are shown relative to the basal control condition, which is set at 1 (dashed line). Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
The star transcript levels were upregulated by PGE2 in the presence of Fsh (Fig. 5C). However, Fsh-stimulated androgen release, being about 10-fold higher than basal (Fig. 4D, left panel), was not changed by PGE2 (Fig. 5D, left panel), suggesting that PGE2 is not acutely modulating basal or gonadotropin-stimulated testicular androgen production. Fsh tended to reduce PGE2 production (Fig. 5D, right panel), but different from the androgen effect (Fig. 4D, right panel), statistical significance was not reached. However, blocking steroid production in the presence of Fsh increased testicular PGE2 release compared to Fsh alone, but not compared to basal conditions, suggesting that this increase reflects the (trilostane-based) removal of steroid-mediated inhibition of PGE2 production. Taken together, the stimulatory effects of both, Fsh and androgens on spermatogonial differentiation were reduced by PGE2, androgens diminished PGE2 production, and PGE2 in combination with androgen induced an accumulation of Aund spermatogonia.
Ep4 receptor mediates PGE2-induced effects in the zebrafish testis
PGE2, but not PGD2 or PGF2α, modulated the activity of spermatogonia. To gain additional knowledge on the mechanism mediating PGE2 effects, we aimed at investigating which receptor(s) mediate these effects. We have found previously (Crespo et al. 2016, Morais et al. 2017) that two PG synthases (ptgs2a and ptges3b) and two receptors (ptger2a and ptger4b) were modulated by different hormones and growth factors (summarized in Fig. 6A). In situ hybridization studies failed to localize the cellular expression of ptger2a and ptger4b, possibly due to their low expression levels in the testis (Table 2). Using alternative approaches, we first, analyzed ptger2a and ptger4b transcript levels available from an RNAseq dataset that compares control, germ cell-depleted, and recovering testes (data set GSE116611). This suggested an enrichment of receptor expression in somatic cells, since ptger2a and ptger4b transcript levels were upregulated in germ cell-depleted testes and returned to control levels during the recovery of spermatogenesis (Fig. 6B). Enrichment of the ptger4b transcript in somatic cells was confirmed using testes from transgenic Tg(vasa:EGFP) zebrafish. Analyzing FACS-sorted testicular cell suspensions by qPCR showed that ptger4b followed an expression pattern similar to the one obtained for Leydig cell marker insl3 (Fig. 6C). This did not apply to the same stringency to ptger2a that followed the Sertoli cell marker gsdf (Fig. 6C) that probably reflects re-association of some EGFP-positive germ cells with Sertoli cells. Therefore, it cannot be excluded that some ptger2a expression was associated with germ cells.
Ep4 receptor (ptger4b) mediates PGE2 signaling in the zebrafish testis. (A) Modulation of PG-related gene expression in testis tissue cultures exposed to different treatments, and analyzed by qPCR or RNAseq approaches. (B) Expression levels of PGE2 receptors in control, germ cell-depleted (by exposure to the cytostatic agent busulfan, as previously described (Nobrega et al. 2010)), and testes with recovering (from busulfan) spermatogenesis, as described in the NCBI GEO database (data set GSE116611). (C) qPCR analysis of EGFP- and EGFP+ cell fractions. A testicular cell suspension was obtained from testes of transgenic Tg(vasa:EGFP) fish and subjected to FACS. EGFP- and EGFP+ cell fractions were collected and RNA was isolated for analysis of germ (GC), Leydig (LC) and Sertoli cell (SC) markers, as well as PGE2 receptors. Data are expressed relative to the EGFP- condition, which is set at 1. (D and E) Effects of an Ep4 receptor antagonist on germ cell development. Testicular explants were cultured for 4 days in the absence or presence of GW627368X (5 μM). Testis tissue was used to quantify BrdU labeling indices (D), and the areas occupied by the type A and B spermatogonia (E). Results are expressed as mean ± s.e.m. (n = 5–8) and asterisks indicate significant differences between groups (*P < 0.05, **P < 0.01, ***P < 0.001). Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0309.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
The Ep4 receptor antagonist GW627368X increased the proliferation activity and relative numbers of differentiating spermatogonia (Fig. 6D and E), a response similar to the one seen when blocking PG production by INDO (Fig. 2A and B). Analyzing the transcript levels also quantified when blocking PG production by INDO (Fig. 2D), we did not find changes in these transcript levels in response to GW627368X (data not shown). Since ptger2a transcript levels were modulated in response to Amh (Fig. 6A) and may be associated with germ cells (Fig. 6C), we also tested the Ep2 receptor antagonist PF04418948. However, this compound did not affect the proliferation activity or relative numbers of spermatogonia (data not shown). Taken together, our results indicate that the b paralogue of the Ep4 receptor is relevant for mediating PGE2 action in the zebrafish testis, and that expression of this receptor is associated preferentially with testicular somatic cells.
Discussion
Our main findings are that locally produced PGE2, but not PGD2 or PGF2α, reduced the proliferation activity of type Adiff and/or type B spermatogonia, both under basal and Fsh- or androgen-stimulated conditions. Moreover, PGE2 increased the self-renewal proliferation of type Aund spermatogonia resulting in their accumulation, which depended on a permissive effect of Fsh-stimulated androgen/steroid productions. Hence, local PGE2 signaling in the testis lowered the production of differentiating spermatogonia but expanded the pool of type Aund spermatogonia, suggesting overall a reduced spermatogenic activity. An inhibitory effect of PGE2 on spermatogenesis was reported earlier based on in vivo studies in mammals (mice (Abbatiello et al. 1975) and dogs (Moskovitz et al. 1987)), describing reduced numbers of spermatocytes and spermatids.
Considering the site of PG production, we found Cox2 protein in Sertoli cells contacting type A spermatogonia. In rat (Winnall et al. 2007) and dog (Korber & Goericke-Pesch 2019) testis, COX2 likewise was present in Sertoli cells but also in Leydig cells, which was not the case in zebrafish. COX2 produces PGH2, which requires PGE synthase to be converted to PGE2. The ptges3b transcript has been localized to Sertoli cells contacting Adiff and B type spermatogonia and spermatocytes in zebrafish (Crespo et al. 2016). This opens the possibility that in tubuli showing a high number of these more differentiated germ cells high PGE2 levels are present, a setting compatible with a local negative feedback loop to reduce the production of further differentiating spermatogonia while expanding the pool of Aund (Fig. 7).
Schematic illustration showing the endocrine and paracrine regulation of zebrafish testicular PGE2 production and the stages of spermatogonial development affected by PGE2. Described effects by secreted factors are indicated by black (Fsh), green (11-KT), red (Amh) and orange (PGE2) arrows, while germ cell development or germ cell-mediated effects are indicated in grey. Fsh, follicle-stimulating hormone; 11-KT, 11-ketotestosterone; Amh, anti-Müllerian hormone; PGE2, prostaglandin E2; Aund, type A undifferentiated spermatogonia; Adiff, type A differentiating spermatogonia; B, type B spermatogonia. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0309.
Citation: Journal of Endocrinology 244, 1; 10.1530/JOE-19-0309
Although zebrafish Leydig cells were negative for Cox2 or ptges3b, Leydig cell-derived androgens are not only stimulators of spermatogenesis (Crowder et al. 2018, de Castro Assis et al. 2018, Tang et al. 2018), but also seem relevant in the context of PG signaling. Part of the stimulatory action of androgens on spermatogenesis may be related to reducing testicular PGE2 production via increasing hpgd mRNA, coding for an enzyme that catabolizes PGs. Also decreasing transcript levels of ptger4b can contribute to an androgen-mediated limitation of PGE2 signaling. Different from androgens, another stimulator of spermatogenesis, Fsh (Crespo et al. 2016) had no rapid (overnight) inhibitory effect on PGE2 production but reduced the transcript levels of two PG synthases (ptgs2a and ptges3b) after 4 days of tissue culture. Therefore, it seems possible that androgen-independent effects of Fsh also reduce PGE2 synthesis. FSH-modulated Cox2 expression and PG production has been reported from primary Sertoli cell cultures of immature rat (Jannini et al. 1994) or adult hamster (Matzkin et al. 2012). It remains to be clarified what the functional connotation is of the nuclear localization of Cox2 in zebrafish Sertoli cell nuclei; a nuclear localization of COX2 has been reported earlier for rat spermatogonia (Neeraja et al. 2003).
While stimulators of spermatogenesis (androgens and Fsh) inhibited PGE2 signaling, we have found previously that Amh, an inhibitor of zebrafish spermatogenesis (Skaar et al. 2011), promoted PGE2 signaling: Amh elevated transcript levels for ptgs2a and ptger4b (Morais et al. 2017), compatible with increasing PG production and PGE2 sensitivity. A schematic working model summarizing effects of regulators of PGE2 production is presented in Fig. 7, placing PGE2 downstream of Fsh and intratesticular factors targeted by Fsh, such as androgens and Amh. However, we did not find evidence for PG effects on Leydig cell androgen production. This is different from the situation in mammals (see ‘Introduction’ section), and also from previous work in a relative of the zebrafish, the goldfish, reporting PGE2-stimulated testicular androgen production (Wade & Van der Kraak 1993). However, later experiments involving treatment of adult zebrafish with an NSAID (ibuprofen) did not result in changes of sex steroid plasma levels (Morthorst et al. 2012). Similarly, in our experiments, PGE2 did not modulate acute (overnight) basal or Fsh-stimulated androgen release but PGE2 did change star, cyp17a1 or ar transcript levels after 3 or 4 days of tissue culture. We may have missed such intermediate-term effects of PGE2 on androgen production with our approach, but the type of effect is not clear. For example, we recorded reduced star and cyp17a1 transcript levels when blocking PG production, but only when steroid production was not blocked at the same time (Figs 2 and 3). One possibility to understand this data set is assuming that PGs may prevent steroid-induced downregulation of star and cyp17a1. However, this possibility does not fit to the PGE2-induced decrease of star, cyp17a1 and ar transcript levels when 11-KT was present in the incubation medium (Fig. 4C). More work is required to clarify the relation between PG and androgen/steroid production. For example, differences between observations made by blocking PG production using INDO and adding PGE2 may reflect effects of PGs other than PGE2 that are also no longer produced when using INDO.
Our working model also summarizes the main effects of PGE2 on type A and B spermatogonia (Fig. 7). We recorded inhibitory effects on the development of type Adiff and B spermatogonia that were independent of the absence or presence of steroids/androgen or Fsh. The self-renewal proliferation of type Aund spermatogonia, on the other hand, was stimulated by PGE2 but this effect depended either on the presence of steroids/androgen (Figs 2, 3 and 4), or on undisturbed steroid production in the presence of Fsh (Fig. 5 of the present MS and Fig. 6 of Morais et al. 2017). It therefore appears that steroids play a (yet to be characterized) permissive role for the PGE2-triggered effects on type Aund spermatogonia. In addition, an unexpected, PG-independent androgen effect on type Aund spermatogonia became evident when comparing the results presented in Figs 2 and 3. In these two experiments, PG production was blocked but it required the additional blocking of steroid production to decrease Aund proliferation and to partially deplete these cells (Fig. 3). This suggested that steroid hormones otherwise support self-renewal proliferation of Aund. Previous work has shown already that PGE2 promoted self-renewal proliferation and accumulation of Aund in the presence of Fsh and undisturbed steroid production (Fig. 6C and D in Morais et al. 2017). In the light of the results presented in Figs 2 and 3, we speculated that this effect seen in response to the combined presence of Fsh and PGE2, may depend on the capacity to produce biologically active steroids. Repeating this experiment in the presence of trilostane indeed abolished the effect of PGE2 on Aund but not on the more differentiated Adiff and B spermatogonia (Fig. 5A and B). Since androgens are likely candidates for mediating this action, we examined if the PGE2 effect on Aund spermatogonia is recovered in the presence of 11-KT, which indeed was the case (Fig. 4). Hence, in the absence of PGs, sex steroids alone already stimulate self-renewal/accumulation of Aund, which is further stimulated when PGE2 is present as well (Fig. 7). Also in salmon testis tissue collected from fish that have just entered pubertal development and showed an elevated level of single cell proliferation activity, PGE2 production is upregulated (Crespo et al. 2019). Since in these males fshb transcript levels in the pituitary and plasma androgen levels were also elevated, it seems possible that in salmon testis, a combined Fsh/androgen/PGE2 signaling accompanies the expansion of the population of type Aund spermatogonia observed in the early pubertal salmon testis (Crespo et al. 2019, Schulz et al. 2019).
Zebrafish testis tissue expresses eight PGE2 receptor subtypes (Table 2). Previous work (Morais et al. 2017) suggested that Ptger2a and Ptger4b are relevant candidates and the present experiments involving EP2 and EP4 receptor antagonists further narrowed it down to Ptger4b. The latter is preferentially expressed by testicular somatic cells, showing in particular a pattern similar to the Leydig cell marker gene insl3. Interestingly, the only growth factor gene that responded to PGE2 with elevated transcript levels was wnt5a (Fig. 4C). Expression of this Wnt ligand is stimulated by Fsh in Leydig cells in the zebrafish testis and stimulates the self-renewal proliferation of type Aund spermatogonia (Safian et al. 2018). Therefore, one possibility for PGE2 to promote the self-renewal may involve elevated wnt5a expression. However, so far, we cannot offer a mechanistic hypothesis for the inhibitory effect of PGE2 on type Adiff and B spermatogonia.
Other PGs had no effect in zebrafish in our experiments. In mice, PGD2 inhibited the differentiation of fetal spermatogonia, using different mechanisms that relied on Sertoli and/or germ cell expression of PGD2 receptors (Moniot et al. 2014, Rossitto et al. 2015). These receptors are not expressed by the adult zebrafish testis (Table 2). Work on bladder cancer stem cells showed that PGE2 promoted stem cell production (Kurtova et al. 2015). Similarly, PGE2 treatment increased hematopoietic stem cell numbers and cyclooxygenase activity was required for their formation (North et al. 2007). Recently, it has also been shown that isolated human testicular peritubular cells secrete PGE2 and that PGE2, as well as EP1 and EP4 receptor agonists, increased glia cell line-derived neurotrophic factor (GDNF) levels (Rey-Ares et al. 2018), a growth factor that maintains the stem cell pool by promoting SSC self-renewal proliferation (Chen et al. 2016, Potter & DeFalco 2017). Evidence for a potential role of Gndf in the rainbow trout testis to prevent differentiation of Aund spermatogonia has been published (Bellaiche et al. 2014). Although a gdnf gene has been annotated in the zebrafish genome, information on its functions in the testis is not available yet.
Taken together, it appears that prostaglandin signaling inhibits the production of more differentiated cells while stimulating stem cell self-renewal in undifferentiated cells in a range of vertebrate models and cell types. For the adult zebrafish testis, we propose that PGE2 signaling is an integral part of the local regulatory network used by reproductive hormones to adjust spermatogenic activity to the changing requirements via endocrine signals, in particular pituitary Fsh, while safeguarding testis tissue homeostasis.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This study was co-funded by the Research Council of Norway BIOTEK2021/HAVBRUK program with the projects SALMAT (n° 226221) and SALMOSTERILE (n° 221648). The authors thank the financial support provided by the National Council for Scientific and Technological Development of Brazil (CNPq; project n°: 202195/2015-5) and the China Scholarship Council (CSC; grant n°: 201706310069), for the scholarships awarded to M S L and Y T Z, respectively.
Acknowledgements
The authors thank Esther Hoekman, Daniëlle Janssen and Henk Westland for maintaining the zebrafish stocks and Henk van de Kant (all from the Science Faculty, Utrecht University, The Netherlands) for technical support regarding histology and immunohistochemistry. Also, they thank Dr Ger J A Arkesteijn (Faculty of Veterinary Medicine, Utrecht, The Netherlands) for expert assistance with the FACS analysis, and Laura Franken and Martijn Breeuwsma (Master students at Utrecht University, The Netherlands) for their help during initial stages of the work.
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