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. Author manuscript; available in PMC: 2016 Apr 1.
Published in final edited form as: Dev Biol. 2015 Jan 23;400(1):33–42. doi: 10.1016/j.ydbio.2015.01.013

Ecdysone response gene E78 controls ovarian germline stem cell niche formation and follicle survival in Drosophila

Elizabeth T Ables 1,3,4,†, Kelly E Bois 1,3, Caroline A Garcia 4, Daniela Drummond-Barbosa 1,2,3,*
PMCID: PMC4448935  NIHMSID: NIHMS658449  PMID: 25624267

Abstract

Nuclear hormone receptors have emerged as important regulators of mammalian and Drosophila adult physiology, affecting such seemingly diverse processes as adipogenesis, carbohydrate metabolism, circadian rhythm, stem cell function, and gamete production. Although nuclear hormone receptors Ecdysone Receptor (EcR) and Ultraspiracle (Usp) have multiple known roles in Drosophila development and regulate key processes during oogenesis, the adult function of the majority of nuclear hormone receptors remains largely undescribed. Ecdysone-induced protein 78C (E78), a nuclear hormone receptor closely related to Drosophila E75 and to mammalian Rev-Erb and Peroxisome Proliferator Activated Receptors, was originally identified as an early ecdysone target; however, it has remained unclear whether E78 significantly contributes to adult physiology or reproductive function. To further explore the biological function of E78 in oogenesis, we used available E78 reporters and created a new E78 loss-of-function allele. We found that E78 is expressed throughout the germline during oogenesis, and is important for proper egg production and for the maternal control of early embryogenesis. We showed that E78 is required during development to establish the somatic germline stem cell (GSC) niche, and that E78 function in the germline promotes the survival of developing follicles. Consistent with its initial discovery as an ecdysone-induced target, we also found significant genetic interactions between E78 and components of the ecdysone signaling pathway. Taken together with the previously described roles of EcR, Usp, and E75, our results suggest that nuclear hormone receptors are critical for the broad transcriptional control of a wide variety of cellular processes during oogenesis.

Keywords: Eip78C, ecdysone signaling, oogenesis, germline stem cell

Introduction

Nuclear hormone receptors play key roles in reproduction (Pestka et al., 2013). The nuclear hormone receptor superfamily consists of seven classes of transcription factors containing a common DNA-binding motif, many of which are regulated by small lipophilic molecules (Evans and Mangelsdorf, 2014). Nuclear hormone receptors control many aspects of female gonad development and function; for example, the classic steroid receptors Progesterone Receptor and Estrogen Receptor regulate follicle development, ovulation, and luteinization, while the orphan nuclear receptor Liver Receptor Homolog-1 is necessary for maintenance of the corpus luteum and formation of the placenta (Pestka et al., 2013; Zhang et al., 2013). Due in large part to the high degree of nuclear hormone receptor gene duplication in vertebrates, however, the function of most of these receptors in reproduction remains unclear.

The nuclear hormone receptor superfamily is well conserved from vertebrates to invertebrates; indeed, Drosophila melanogaster has 18 nuclear hormone receptors, representing each of the major vertebrate receptor subfamilies, with relatively little gene duplication (King-Jones and Thummel, 2005). Further, despite obvious differences in female reproductive strategies, Drosophila and mammalian oogenesis share a wide variety of common characteristics (Matova and Cooley, 2001; Pepling et al., 1999; Sun and Spradling, 2013). Drosophila ovaries are composed of 15 to 20 ovarioles, or strings of progressively older follicles that ultimately develop into a mature oocyte (Fig. 1A). Oogenesis is fueled by the activity of germline stem cells (GSCs), which reside in a somatic niche at the anterior of each ovariole (within a structure called the germarium) and produce daughter cells via asymmetric divisions. The GSC progeny undergo four mitotic divisions with incomplete cytokinesis to form 16-cell cysts, each comprised of the oocyte plus 15 supporting nurse cells. Each germline cyst is encapsulated by somatic follicle cells, forming a follicle that will progress through 14 developmental stages prior to egg deposition. Nuclear hormone receptor signaling regulates oogenesis in both Drosophila and mammals (Belles and Piulachs, 2014; Pestka et al., 2013); therefore, understanding the role of nuclear hormone receptors in the Drosophila ovary may yield important insights into mammalian reproduction.

Fig. 1.

Fig. 1

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E78 is expressed in the ovarian germline. (A) The Drosophila ovary is made of 15–20 ovarioles, each harboring a germarium (G; top) and progressively older follicles. Within the germarium, germline stem cells (GSCs; green) are juxtaposed to cap cells, the major cellular component of the somatic niche (yellow), and a subset of escort cells (gray). GSCs divide to form daughter cells, which divide four additional times to form 16-cell germline cysts (green) composed of nurse cells (nc) and an oocyte (oo). Follicle cells (fc; red) encapsulate each cyst, forming a follicle that pinches off from the germarium and progresses through 14 stages of oogenesis. (B) The E78 genomic locus encodes four mRNA isoforms corresponding to two distinct proteins; RA, RD, and RE produce a nuclear hormone receptor with a canonical DNA-binding domain (DBD) and ligand-binding domain (LBD), while RB codes for a significantly shorter protein that lacks a DBD. Red arrowheads represent transposon insertions used in this study, dotted red lines indicate affected regions in the E78Δ31 and E788h alleles, and solid red lines show regions targeted by RNAi transgenes. Thin black arrows indicate the locations of RT-PCR primers. (C) E78::GFP ovariole labeled with anti-GFP (green), anti-Hts (red; fusomes and follicle cell membranes), and anti-LamC (red; nuclear envelope of cap cells). Arrows, nurse cells; arrowheads, germ cells in Region 2b/3. (D) X-gal detection of β-gal expression in hsGal4-E78-LBD/+;UASp-lacZ/+ ovariole. Scale bars, 20 μm. (E) RT-PCR for E78, in comparison to a ubiquitously expressed control transcript (Rp49), demonstrates a significant reduction in E78 mRNA levels in E78Δ31/Df mutants, yet only a modest reduction in E788h/Df mutants.

Nuclear hormone receptors regulate many aspects of Drosophila oogenesis. The steroid hormone ecdysone is produced by late stage ovarian follicles (Huang et al., 2008) and binds to a heterodimeric receptor consisting of two nuclear hormone receptors, Ecdysone Receptor (EcR) and Ultraspiracle (Usp), both of which are widely expressed in the ovary (Riddiford et al., 2000). Ecdysone signaling is required for ovary development prior to adulthood (Gancz et al., 2011), as well as for the proliferation and self-renewal of adult GSCs (Ables and Drummond-Barbosa, 2010). Germline cyst formation and encapsulation are indirectly dependent on ecdysone signaling (Konig et al., 2011; Morris and Spradling, 2012). Ecdysone signaling also controls follicle growth and development (Buszczak et al., 1999; Carney and Bender, 2000), as well as the migration of border cells, a subpopulation of follicle cells (Bai et al., 2000; Jang et al., 2009). Other nuclear hormone receptors also have known roles in oogenesis. For example, Hormone receptor 39 (Hr39) is required for development of the reproductive tract (Allen and Spradling, 2008; Sun and Spradling, 2012), while Ecdysone-induced protein 75B (E75) controls a variety of cellular processes in the ovary, including follicle survival during early vitellogenesis (Buszczak et al., 1999; Morris and Spradling, 2012). The function of most other Drosophila nuclear hormone receptors in oogenesis, however, remains unexplored.

The nuclear hormone receptor Ecdysone-induced protein 78C (E78), which is closely related to E75 and to mammalian Rev-Erb and Peroxisome Proliferator Activated Receptors (Bridgham et al., 2010; King-Jones and Thummel, 2005), was also originally identified as an early ecdysone target (Stone and Thummel, 1993). Analysis of a homozygous viable hypomorphic E78 mutant, however, revealed few critical physiological functions, and no overt defects in female fertility (Russell et al., 1996). In this study, we performed a comprehensive analysis of the function of E78 during Drosophila oogenesis using targeted deletion of the E78 locus and germline-enhanced RNAi-mediated knockdown. Remarkably, we found that loss of E78 causes multiple ovarian defects, resulting in significantly reduced female fertility. Specifically, E78 is required for the establishment of the proper number of GSCs during development, and for follicle viability in adult females. In addition, we demonstrated a maternal requirement for E78 during early embryogenesis. Consistent with its original identification as a target of ecdysone signaling, we also find that E78 genetically interacts with EcR and ecdysoneless (ecd) to control follicle survival. Taken together, our work identifies novel roles for E78 in Drosophila, and suggests that E78 may be an important mediator of the ecdysone-induced transcriptional cascade in oogenesis.

Results

E78 is widely expressed in the ovarian germline

The E78 gene locus encodes four mRNAs corresponding to two proteins that include a common ligand-binding domain; E78-RA/RD/RE also has a canonical DNA-binding domain characteristic of nuclear hormone receptors, whereas E78-RB lacks a DNA-binding domain (Fig. 1B). To assess E78 localization in the ovary, we took advantage of two E78 reporters. E78::GFP carries a bacterial artificial chromosome (CH321-05P13) in which the native E78 promoter drives the expression of an E78 fusion protein with a carboxy-terminal GFP tag (www.flybase.org). We found E78::GFP expression in nurse cell nuclei beginning in region 2b/3 of the germarium through stage 10 of oogenesis (Fig. 1C and Fig. S1). We noted a similar pattern of expression using the ligand-sensor line hsGal4-E78-LBD, in which the E78 ligand-binding domain is fused to a heat-shock inducible Gal4 (Palanker et al., 2006). Using the germline-compatible UASp-lacZ reporter to detect activation of the ligand sensor, we observed E78 activity in the germline, particularly in stages 3 through 8 of oogenesis (Fig. 1D and Fig. S1I–J). We also detected somatic expression of both E78 reporters in stages 8 to 12 follicles, both in main body follicle cells and pre- and post-migratory border cells (Fig. S1D–J). Taken together, these data suggest that the E78 receptor and its as-yet-unidentified ligand are present in the Drosophila ovary.

E78 is required for female fertility

Although previous analysis of a hypomorphic inversion allele (E788h) failed to find obvious phenotypes in the ovary (Russell et al., 1996), the detection of E78 reporters in the germline prompted us to create a new E78 mutant allele. We used Flippase (FLP)-mediated recombination between two available PiggyBac insertions carrying Flippase recognition target (FRT) sequences to remove the translational start site of all E78 isoforms and the DNA-binding domain of isoforms RA/RD/RE (Fig. 1B and Fig. S2), thereby generating the E78Δ31 allele. When placed in trans to Df(3L)BSC419, a deficiency that uncovers the entire E78 gene locus (www.flybase.org), we found that E78Δ31 mutants display greatly reduced levels of E78 mRNA (Fig. 1E). Although the precise molecular nature of the E788h allele is unclear, RT-PCR demonstrates that, in contrast to E78Δ31 mutants, E78 mRNA is still robustly produced in E788h/Df(3L)BSC419 mutants. We therefore focused our phenotypic analysis on the E78Δ31 allele. Initial screening indicated that E78Δ31 mutants were homozygous viable and male fertile, yet completely female sterile. As a potentially interesting aside, we serendipitously noticed that E78Δ31 mutants failed to eclose at Mendelian frequencies when raised on a non-standard Drosophila diet high in sugar (Table S1); lethality on this diet was rescued by the addition of yeast paste to cultures.

To further investigate the cause of the female sterility in E78Δ31 mutant females, we quantified egg production in E78 mutant females at different ages (Fig. 2A). Consistent with their abundant levels of E78 mRNA, E788h hemizygous females laid similar numbers of eggs as their control siblings. In contrast, E78Δ31 hemizygous females produced roughly one-third as many eggs as their wild-type sibling controls; egg production continued to decline with time, such that E78Δ31 mutant females were virtually sterile by two weeks post-eclosion.

Fig. 2.

Fig. 2

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E78 is required for optimal female fertility. (A) Five pairs of E788h/Df and E78Δ31/Df mutant and heterozygous sibling control (E788h or Df/TM3, and E78Δ31 or Df/TM3, respectively) females per bottle (in triplicate) were kept on wet yeast paste beginning one day after eclosion, and the number of eggs laid per female was quantified at three, 6, 9, 12, and 14 days after eclosion. Error bars, mean ± SEM. (B) Fifty eggs laid by mutant and control females at three days or 14 days after eclosion (in triplicate) were monitored for hatching. *p < 0.01, ***p < 0.0001; Student’s two-tailed T-test.

Because the reduction in egg laying by E78Δ31 hemizygous females alone could not sufficiently explain the complete sterility observed in our initial fertility assay, we asked whether there might also be a maternal requirement for E78 function (Fig. 2B). We found that the number of viable embryos (quantified as the percentage of embryos that hatched) produced by E78Δ31 hemizygous mothers was significantly decreased relative to that of control females; this effect was more pronounced in two-week-old E78Δ31 mutant mothers than three-day-old mothers. Maternal E78-deficient embryos progressed beyond fertilization, as evidenced by the normal completion of the first few embryonic syncytial divisions, but the majority of embryos derived from E78Δ31 females failed to gastrulate (E.T.A. and D.D.B., unpublished observations). Interestingly, although E788h mutant females displayed no defects in the number of eggs laid, the fraction of viable embryos produced by these females was also significantly decreased, suggesting that insufficient levels of maternal E78 mRNA are present in their early embryos. Taken together, these results indicate that E78 is required for proper female fertility due to its roles during oogenesis for adequate egg production levels, and in the early embryo for progression through development.

E78 is not required for adult germline stem cell proliferation or maintenance

We next sought to understand the cellular mechanisms underlying the dramatic decrease in egg production in E78Δ31 hemizygous females. Lower rates of GSC proliferation might phenotypically manifest in females as decreased egg production. GSCs and their progeny can be easily identified by co-immunofluorescence detection of Vasa, a germ cell-specific protein, and Hts, a component of the fusome, a germline-specific organelle (Lasko and Ashburner, 1990; Lin et al., 1994). As a measure of GSC proliferation rate, we calculated the percentage of GSCs in S-phase, based on incorporation of the thymidine analog 5-ethynyl-2′-deoxyuridine (EdU) during a pulse immediately following dissection, as described (Ables and Drummond-Barbosa, 2013). There was no significant reduction in GSC proliferation in E78Δ31 mutant GSCs (8.8% EdU-positive GSCs; n = 250 GSCs) compared to control GSCs (9.1% EdU-positive GSCs; n = 287 GSCs), indicating that decreased GSC proliferation is not the underlying cause of the decreased egg production observed in E78Δ31 mutants.

Although E78Δ31 mutant females have normal rates of GSC proliferation and normal overall germarium morphology, we noticed that the germaria of E78Δ31 hemizygous females were significantly smaller than those of heterozygous sibling controls (Fig. 3A,B). We therefore hypothesized that there could be differences in the number of GSCs, and thus, the number of progeny in E78Δ31 mutant germaria. We quantified GSC number in E78Δ31 mutant females (Fig. 3C–D) by counting Vasa-positive cells with an anteriorly-localized fusome juxtaposed to cap cells (a component of the somatic niche; labeled with the nuclear membrane marker Lamin C), as described (Ables and Drummond-Barbosa, 2010). In newly eclosed females, we noted a small, yet significant decrease in GSC number, as compared to wild-type sibling controls: the average number of GSCs per germarium was decreased (Fig. 3C), and the proportion of germaria with only one GSC was increased in E78Δ31 mutants (Fig. 3D). These results suggest a developmental role for E78 in establishing proper GSC numbers.

Fig. 3.

Fig. 3

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E78 is required for establishing the proper number of GSCs. (A–B) Heterozygous sibling control (A) or E78Δ31/Df mutant (B) germaria labeled with anti-Vasa (green; germ cells), anti-Hts (red; fusomes and follicle cell membranes), and anti-LamC (red; nuclear envelope of cap cells). Asterisks, cap cells. Scale bar, 5 μm. (C) Average number of GSCs per germarium in heterozygous sibling control or E78Δ31/Df hemizygous females at 0–2 days, 1 week, 2 weeks, or 3 weeks after eclosion. The number of germaria analyzed is shown inside bars. Error bars, mean ± SEM. ***p < 0.0001; Student’s two-tailed T-test. (D) Frequencies of germaria containing 0 (red), 1 (white), 2 (gray), or 3 or more (black) GSCs per germarium.

To assess whether E78Δ31 mutant GSCs have maintenance defects, we counted GSCs in adult females over time. E78Δ31 hemizygous females and their sibling controls exhibited very similar rates of decline in GSC number over time (Fig. 3C,D), indicating that E78 is not required for GSC maintenance in adult females. We also used FLP/FRT-mediated mitotic recombination to generate E78Δ31 homozygous mutant GSC clones in the context of wild-type tissue. Mutant GSCs and their progeny were recognized by the absence of a β-galactosidase (β-gal) marker, and germaria containing β-gal-negative GSC progeny, but no β-gal-negative mother GSC were scored as a GSC loss event (Ables and Drummond-Barbosa, 2010). Mock (in which both β-gal-negative and -positive cells are wild-type) and E78Δ31germline mosaic germaria had comparable levels of GSC loss [24% GSC loss in mock mosaics (n = 25 germaria) versus 25% GSC loss in E78Δ31 mosaics (n = 48 germaria)], indicating that even in the presence of neighboring wild-type GSCs, E78Δ31 GSCs are well maintained. Taken together, these data show that E78 is required during ovary development for the establishment of proper number of GSCs, but not for adult GSC maintenance.

E78 is required in ovarian somatic precursor cells to establish proper GSC niche size

To test if the decreased number of GSCs in newly eclosed E78Δ31 mutant germaria was due to a requirement for E78 in germ cells during development for the establishment of proper GSC numbers, we used short hairpin interfering RNA (RNAi) to specifically reduce E78 function in developing germ cells via the UAS/Gal4 system (Ni et al., 2011). We generated three germline-enhanced RNAi lines targeting E78: UAS-E78shRNA-A (VALIUM20) and UAS-E78shRNA-B (VALIUM22) target the same sequence in the region corresponding to the ligand binding domain, while UAS-E78shRNA-C (VALIUM22) targets the last exon common to all E78 isoforms (Fig. 1A; see Materials and Methods). We then used the germline-specific driver nos-Gal4::VP16 (Van Doren et al., 1998) to induce E78 knockdown in germ cells at their earliest stages of development, and counted the number of GSCs per germarium in adult females at one week after eclosion (Fig. 4A). We found no significant decrease in the average number of GSCs for any of the RNAi lines tested (which, based on later experiments in this study, are effective), suggesting that E78 might not be required in the developing germline to establish the proper numbers of adult GSCs.

Fig. 4.

Fig. 4

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E78 is required in the developing somatic niche to establish the proper number of GSCs. (A) Average number of GSCs per germarium in nos-Gal4::VP16 control or nos-Gal4::VP16/UAS-E78shRNA knockdown females. (B) Box and whisker plot showing number of cap cells per germarium in heterozygous sibling control or E78Δ31/Df mutant females at 0–2 days, 1 week, 2 weeks, or 3 weeks after eclosion. (C) Number of cap cells per germarium in bab1-Gal4 control or bab1-Gal4/UAS-E78shRNA knockdown females. (D) Average number of GSCs per germarium in bab1-Gal4 control or bab1-Gal4/UAS-E78shRNA knockdown females. The number of germaria analyzed is shown inside (A,D) or below (B,C) bars. Error bars, mean ± SEM. *p < 0.01; **p < 0.0005; ***p < 0.0001; n.s., not significant; §p < 0.0001, as compared to both control and rescue; Student’s two-tailed T-test.

It is well known that the number of cap cells, which are the major cellular components of the GSC niche, is an important determinant of GSC number (Xie, 2013). We therefore hypothesized that the reduced number of GSCs in E78Δ31mutant germaria could be due to a smaller niche size (i.e. fewer cap cells). Using immunofluorescence for the nuclear envelope marker Lamin C to highlight individual cap cells (Fig. 3A,B), we quantified the number of cap cells per germarium in control and E78Δ31 mutant ovaries (Fig. 4B), and found a small but significant reduction in the average number of cap cells per germarium in E78Δ31 mutants, regardless of age. Thus, smaller niche sizes in E78Δ31 ovaries correlate with the decreased GSC numbers.

To determine whether E78 is required within cap cell precursors to control niche size, we used RNAi to knock down E78 specifically in those cells (Fig. 4C,D). The bab1-Gal4 driver, which is highly expressed in somatic precursors in the developing ovary (Gancz et al., 2011), was used to drive two independent RNAi transgenes (UAS-E78JF02258 and UAS-E78shRNA-A), both of which target sequences common to all four E78 isoforms. We observed a small but significant reduction in cap cell number (Fig. 4C), and a corresponding decrease in GSC number (Fig. 4D), when either RNAi transgene was used to knock down E78 function in the developing niche. These results suggest that E78 is cell autonomously required for generation of the proper number of cap cells and, thereby, to establish normal numbers of GSCs prior to adulthood. Interestingly, introducing one copy of the E78::GFP transgene rescues the number of cap cells, but is not sufficient to rescue GSC numbers, in E78Δ31 females (Fig. S3A–C), suggesting that additional changes in the niche (besides cap cell number) might contribute to GSC loss.

E78 is required for cyst viability during follicle development

The relatively modest reduction in GSCs and cap cells observed in germaria of young E78Δ31 females is not sufficient to explain the drastic and progressive decrease in egg production in these mutants. Following the last germline division, encapsulation of germline cysts by a monolayer of somatic follicle cells produces individual follicles, which bud from the germarium and progress through 14 distinct stages of growth and development prior to egg deposition (Fig. 1A) (Spradling, 1993). Each wild-type ovariole thus contains a series of follicles at progressively more advanced stages of development (Fig. 5A). Abnormal follicle growth, development, or death can result in decreased female fertility; therefore, we analyzed developing E78Δ31 follicles for defects. Early stage E78Δ31 mutant follicles (stages 1–4) of normal morphology, size, and nurse cell chromatin compaction were easily identified (Fig. 5B,C). Intriguingly, we observed a significant number of ovarioles with dying follicles characterized by shrunken appearance and pyknotic or absent nurse cell nuclei (arrows, Fig. 5B,C; quantified in G). Dying follicles were found among normal early follicles (around stages 4–5; Fig. 5B) and late follicles (stages 11–14; Fig. 5C). In some cases, the somatic follicle cell monolayer remained largely intact (Fig. 5B). Introduction of a copy of E78::GFP rescued the follicle death phenotype of E78Δ31 hemizygous mutants (Fig. S3D). In addition, similar phenotypes were observed upon RNAi-mediated knockdown of E78 using the nos-Gal4::VP16 driver (Van Doren et al., 1998), which is strongly expressed in the germline at all stages of oogenesis (Fig. 5E,F; quantified in G). Taken together, these results suggest that E78 is required in the germline to promote follicle survival during oogenesis.

Fig. 5.

Fig. 5

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E78 is required in the germline for follicle survival. (A–C) Heterozygous sibling control (A) or E78Δ31/Df mutant (B, C) ovarioles labeled with anti-Vasa (green; germ cells). (D) E78Δ31 mutant germline mosaic ovariole labeled with anti-β-gal (green; wild-type cells). (E,F) nos-Gal4::VP16/UAS-E78shRNA ovarioles. Anti-Hts (red; fusomes and follicle cell membranes), anti-LamC (red; nuclear envelope of cap cells), and DAPI (blue; nuclei). Arrows indicate dying follicles; asterisks denote intact or partially intact follicle cell monolayer in dying follicles. Scale bars, 50 μm. Oogenesis stages determined (except in the case of dying follicles) by follicle size and nurse cell nuclear morphology, as described (Spradling, 1993). (G–H) Percentage of ovarioles (G) or germline-mosaic ovarioles (H) with at least one dying follicle. (I) Percentage of germline-mosaic ovarioles with at least one cleaved Dcp-1-positive cyst. The numbers of ovarioles (G) or germline-mosaic ovarioles (H, I) analyzed are shown inside bars.*p < 0.05, **p < 0.01, ***p < 0.0001; Chi-square test.

To test whether E78 is cell autonomously required in germline cysts for their survival, we used the FLP/FRT system to create E78Δ31 homozygous germline clones in mosaic ovarioles (Fig. 5D,H and Fig. 6). In stark contrast to mock germline mosaic ovarioles, about 40% of E78Δ31 germline-mosaic ovarioles contained at least one dying follicle, recognizable either by morphology (Fig. 5D,H) or by the expression of cleaved Dcp-1 (Fig. 5I and Fig. S4; McCall and Steller, 1998; Hou et al., 2008), suggesting that E78 mutant cysts die by an apoptotic or autophagic mechanism. Consistent with our observations in global E78Δ31 mutant ovarioles, we noted dying cysts both prior to vitellogenesis (Fig. 6A) and after vitellogenesis (Fig. 6D). We also noted that stage 4/5 follicles containing an E78Δ31 mutant cyst were more frequently observed than follicles at any other stage (stage 6–10; Fig. 6E), suggesting that follicle development may be partially blocked at stage 4/5. Indeed, we noted a decreased number of stage 6–10 follicles containing an E78Δ31 mutant cyst, as compared to mock controls (Fig. 6E). Furthermore, we also occasionally observed E78Δ31 mutant cysts in follicles with a characteristic stage 4/5 morphology, but located posterior to older wild-type follicles (Fig. 6C; 6/107 germline mosaic ovarioles analyzed). Our observations suggest that E78Δ31 mutant cysts are blocked at the stage 4/5 transition, and that a significant number of those that can escape this transition subsequently degenerate.

Fig. 6.

Fig. 6

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Oogenesis stages 4/5 are particularly sensitive to loss of E78 from the germline. (A–D) E78Δ31 germline mosaic ovarioles labeled with anti-β-gal (green; wildtype cells), anti-Hts (red; fusomes and follicle cell membranes), anti-LamC (red; nuclear envelope of cap cells), and DAPI (blue; nuclei). Arrows indicate dying follicles. Scale bars, 20 μm (A–C) or 50 μm (D). Oogenesis stages were determined (where possible) by follicle size and nurse cell nuclear morphology, as described (Spradling, 1993). (E) Distribution of follicles with a β-gal-negative cyst in control or E78Δ31 germline mosaic ovarioles. In wild-type (mock mosaic) ovarioles, follicles containing β-gal-negative cysts at all stages of oogenesis are well represented. In contrast, stages 4/5 follicles with a β-gal-negative cyst are detected at higher frequencies than any other stage in E78Δ31 mutant mosaic ovarioles, while stages 6/7 or 8–10 follicles with a β-gal-negative cyst are under-represented. 85 mock germline mosaic and 107 E78Δ31 germline mosaic ovarioles were analyzed. *p ≤ 0.01; Chi-square test.

E78 genetically interacts with ecdysone signaling to control cyst viability

Like E78, other ecdysone early-response targets, including EcR, E74, and E75, are also required in the germline for follicle viability (Ables and Drummond-Barbosa, 2010; Buszczak et al., 1999; Carney and Bender, 2000). We therefore asked whether E78 functions together with other ecdysone-response genes to control follicle survival (Table 1). Since E78 is remarkably similar in structure to E75, we first tested whether E78Δ31 genetically interacts with a previously described null allele of E75 (Bialecki et al., 2002). Single heterozygous females for E78Δ31 or E75Δ51 displayed minor levels of follicle death (scored as the percentage of ovarioles with at least one dying follicle), and only a small, statistically insignificant increase was observed in double heterozygous females (Table 1). Despite the conservation in protein structure, therefore, we did not find strong genetic interactions between E78Δ31 and E75Δ51, possibly reflecting separate roles for E78 and E75 in the ovary. We then asked whether E78Δ31 genetically interacts with ecdysoneless (ecd), a gene required for the maintenance of proper ecdysone levels (Garen et al., 1977), or EcR, required for reception of the ecdysone signal (Carney and Bender, 2000). As expected, negligible follicle death was observed in ecd1 or EcRM554fs single heterozygous ovaries (Table 1). Remarkably, we found a two-fold increase in follicle death in both ecd1 +/+ E78Δ31 and EcRM554fs/+; E78Δ31/+ double heterozygous females (Table 1). These results suggest that E78 functions together with the ecdysone signaling pathway to control follicle survival, and might reflect an important role for E78 in mediating the ecdysone response during oogenesis.

Table 1.

Genetic interactions between E78Δ31 and ecdysone signaling pathway components.

Genotype % Ovarioles with a dying follicle Number of ovarioles scored p-value
ecd1/+ OR E78Δ31/+ sibling controls 5.14 214
ecd1 +/+ E78Δ31 10.13 227 0.0495*
EcRM554fs/+ OR E78Δ31/+ sibling controls 6.11 229
EcRM554fs/+; E78Δ31/+ 14.81 270 0.0018**
E75Δ51/+ OR E78Δ31/+ sibling controls 7.88 241
E75Δ51 +/+ E78Δ31 11.21 232 0.2182
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*

p < 0.05,

**

p < 0.005 versus sibling controls; Chi-square test.

Discussion

Although nuclear hormone receptors are known to play important roles in a wide variety of biological processes, it remains largely unknown whether or how most of the Drosophila nuclear hormone receptors function during oogenesis. Our study adds to a growing body of literature demonstrating that nuclear hormone receptors are integral to reproductive function at multiple levels, including reproductive organ development, stem cell function, and gamete development and survival (Ables and Drummond-Barbosa, 2010; Buszczak et al., 1999; Carney and Bender, 2000; Evans and Mangelsdorf, 2014; Gancz et al., 2011; Morris and Spradling, 2012; Park-Sarge and Mayo, 1994; Sun and Spradling, 2012, 2013; Zhang et al., 2013). While it remains unclear how E78 contributes mechanistically to the ecdysone signaling network in the ovary, our studies also highlight the intricate connections between Drosophila nuclear hormone receptors and ecdysone signaling. Given the level of structural and functional conservation between Drosophila and mammalian hormonal signaling pathways (King-Jones and Thummel, 2005), we propose that similar connections may exist among diverse mammalian nuclear hormone receptor subtypes. Further studies will be necessary to fully elucidate the molecular networks that tie these pathways together to achieve such important biological regulation.

It is interesting to note that each of the ecdysone early response genes studied in the ovary to date (EcR, E74, E75, E78) encode at least two different mRNA isoforms: one long mRNA isoform resulting from splicing of a very long intron separating conserved DNA- and ligand-binding domains, and a shorter isoform that may or may not produce a distinct protein isoform (www.flybase.org). Previous studies have indicated that ftz-f1, another ecdysone-regulated nuclear hormone receptor, is also encoded by two different mRNA isoforms: ftz-f1-RA (short isoform) is maternally deposited and required for embryogenesis, while ftz-f1-RB (long isoform) is required at other developmental stages (King-Jones and Thummel, 2005). Future studies investigating whether the various isoforms of ecdysone early-response genes differentially control oogenesis versus embryogenesis will help to refine our understanding of how a steroid hormone may induce temporal-, developmental-, and cell type-specific effects.

While our studies demonstrate a specific requirement for E78 in promoting the survival of germline cysts, the mechanisms by which E78 controls cyst survival remain a topic for further exploration. Indeed, mutants of several ecdysone early-response genes, including EcR, E74, and E75, display similar cyst death near the onset of oocyte vitellogenesis (Buszczak et al., 1999; Carney and Bender, 2000), suggesting that ecdysone signaling promotes a maturation or survival cue during follicle development. Very little is known, however, about the targets of ecdysone signaling during earlier previtellogenic stages. Two recent large-scale screens for regulators of ecdysone-regulated cell death in a haemocyte cell line and in salivary glands (Chittaranjan et al., 2009; Wang et al., 2008) may prove useful for identifying targets involved in the decision between cell death and survival. Interestingly, the stage 4/5 cyst death we observed in E78Δ31 mutants is phenocopied by mutations in insulin and target of rapamycin (TOR) signaling pathway components, including InR, chico, TOR, and S6 kinase (Pritchett and McCall, 2012). Since EcR and E78 appear to functionally cooperate, future studies should test whether EcR and E78 regulate members of the insulin/TOR signaling pathways (or vice-versa) to control cyst viability. These basic studies will not only help elucidate the mechanisms by which nuclear hormone receptors control biological processes, but may also add to our general understanding of how nuclear hormone receptor signaling is integrated into other endocrine networks to coordinate cell-specific responses with whole-animal physiology.

Materials and methods

Drosophila strains and culture

Flies were maintained at 22°–25°C in standard molasses medium or in K12 Drosophila diet (U.S. Biologicals) supplemented with yeast paste unless otherwise indicated. For assessment of E78 expression in the ovary, we used a GFP fusion protein, encoded by PBac[Eip78C-GFP.FLAG]VK00037 (E78::GFP; Bloomington Stock Center). For detection of E78 activity, we used the previously described ligand sensor system (Palanker et al., 2006) by crossing P[hsGal4-E78](12.7); P[UAS-eGFP(nls)] (H. Krause) to UASp-lacZ (to detect germline activity) or UASt-nGFP (to detect somatic activity). Females were collected 2–3 days after eclosion and heat-shocked for 1 hour at 37°C, then maintained on wet yeast paste at 29°C for 4 days prior to ovary dissection. For genetic interaction analyses, the following alleles were used: ecd1 (Garen et al., 1977); EcRM554fs (Carney and Bender, 2000); and E75Δ51 (Bialecki et al., 2002). Controls (single heterozygotes carrying a balancer) were compared to double heterozygotes. Balancer chromosomes and other genetic tools are described in FlyBase (www.flybase.org).

Generation of E78 mutants

E78Δ31 mutants were generated via Flippase (Flp)-Flippase Recognition Target (FRT)-mediated excision (Bellen et al., 2011; Parks et al., 2004; Thibault et al., 2004) of approximately 31 kB of E78 genomic sequence, including the predicted translational start site and DNA-binding domain of E78-RA, flanked by the transposable piggyBac insertions PBac[RB]e02923 and PBac[RB]e02853 (Exelixis Collection, Harvard Medical School). Mutant progeny were screened by two-sided PCR (see Fig. S2 and Table S2) (Parks et al., 2004). E78Δ31 mutants were analyzed in trans to Df(3L)BSC419 (Bloomington Stock Center), which uncovers the entire E78 locus. For rescue experiments, we recombined E78::GFP with Df(3L)BSC419, and crossed the resulting line with E78Δ31.

Generation of RNAi lines

UAS-E78shRNA transgenic lines were generated as described (Ni et al., 2008; Ni et al., 2011) (www.flyrnai.org/TRiP-HOME.html). Primers were designed against the carboxy-terminus of E78 (common to all four isoforms) via the Designer of Small Interfering RNA website (http://biodev.extra.cea.fr/DSIR/DSIR.html), using default settings (21 nt siRNA; score threshold 90). Two sequences with corrected scores greater than 85 and no predicted off-targets were selected for hairpin generation: E7826: passenger strand, GATTGTGGAGTTTGCGAAACG; guide strand, CGTTTCGCAAACTCCACAATC; E7846: passenger strand, CGGGCTGAGTGACACCGAAAT; guide strand, ATTTCGGTGTCACTCAGCCCG. Primers were designed according to TRiP recommendations, annealed to form double-stranded oligos, and ligated into EcoRI/NheI-digested pVALIUM20 or pVALIUM22 (Ni et al., 2011), resulting in three UAS-E78shRNA vectors: pUAS-E78shRNA-A (pVALIUM20) and pUAS-E78shRNA-B (pVALIUM22) target E78-RA exon 5 (sequence 26 above), while pUAS-E78shRNA-C (pVALIUM22) targets E78-RA exon 6 (sequence 46 above). Transgenic flies were generated by inserting the pUAS-E78shRNA constructs via the phiC31 site-specific integrase into the attP2 site on the third chromosome (Genetic Services). UAS-E78shRNA-A and P[TRiP.JF02258]attP2 (pVALIUM10, Bloomington Stock Center; targets E78-RA exon 6) were crossed to bab1-Gal4 (Bolivar et al., 2006) to reduce E78 levels in the niche. UAS-E78shRNA-B and UAS-E78shRNA-C were crossed to nos-Gal4 (nos-GAL4::VP16-nos.UTR) to reduce E78 levels in the germline. For both analyses, female progeny were collected 2–3 days after eclosion and maintained for 5 days at 29°C on wet yeast paste prior to ovary dissection by standard protocols.

Clonal analysis

For FLP/FRT-mediated genetic mosaic analyses (Xu and Rubin, 1993), E78Δ31 was recombined with P[neoFRT]80B using standard crosses. Other genetic tools are described in FlyBase. Genetic mosaics were generated by FLP/FRT-mediated recombination in 2–3-day old females carrying a mutant allele in trans to a wild-type allele (linked to an armZ marker) on homologous FRT arms, and a hs-FLP transgene, as described (Ables and Drummond-Barbosa, 2010). A wild-type FRT80B chromosome without armZ was used in lieu of E78Δ31 FRT80B for control mosaics. GSCs were identified based on the juxtaposition of their fusomes to the interface with adjacent cap cells (Ables and Drummond-Barbosa, 2010). GSC loss was measured as the percentage of total germline-mosaic germaria showing evidence of recent GSC loss; namely, the presence of β-gal-negative cystoblasts/cysts in the absence of the β-gal -negative mother GSC, at 6 days after recombination, as described (Ables and Drummond-Barbosa, 2010).

Immunofluorescence, X-gal staining, and microscopy

Ovaries were dissected, fixed, washed, and blocked as described (Ables and Drummond-Barbosa, 2010), except as noted below. The following primary antibodies were used overnight at 4°C: mouse anti-Hts [1B1, Developmental Studies Hybridoma Bank (DSHB); 1:10], mouse anti-Lamin C (LamC) (LC28.26, DSHB; 1:100), rabbit anti-Vasa (a gift from P. Lasko; 1:1000; (Lasko and Ashburner, 1990), rat anti-Vasa (DSHB; 1:50), chicken anti-βgal (ab9361, Abcam; 1:2000), rabbit anti-GFP (TP401, Torrey Pines; 1:2500), and rabbit anti-cleaved Dcp-1 (Cell Signaling #9578; 1:100). For anti-β-gal labeling, fixed ovaries were permeabilized for 30 minutes in 0.5% Triton X-100 in phosphate-buffered saline (PBS; 0.01M phosphate buffer, 0.0027M KCl, and 0.137M NaCl, pH 7.4) prior to blocking. Following a two-hour incubation with Alexa Fluor 488-, 568-, or 633-conjugated goat species-specific secondary antibodies (Life Technologies; 1:200), ovaries were stained with 0.5 μg/ml 4′-6-diamidino-2-phenylindole (DAPI) (Sigma). Ovaries were mounted in 90% glycerol containing 20 mg/ml n-propyl gallate (Sigma). Confocal Z-stacks (1 μm optical sections) were collected with a Zeiss LSM 700 microscope using ZEN Black 2011 software. Images were analyzed and minimally and equally enhanced via histogram using Zeiss ZEN software. Statistical analyses were performed using the Chi-square test, Student’s two-tailed t-test, or one-way ANOVA in Microsoft Excel or GraphPad Prism.

X-gal detection of β-gal activity was performed as described (Margolis and Spradling, 1995), with minor modifications. In brief, ovaries were dissected and teased apart in Grace’s Insect Media (Lonza), then fixed for 8 minutes in 0.5% glutaraldehyde diluted in Grace’s media. Following a 10 minute wash in 0.1% Triton X-100 in PBS, ovaries were incubated overnight at room temperature in X-gal staining solution {10 mM NaH2PO4-Na2HPO4 (pH 7.2), 150 mM NaCl, 1 mM MgCl2·6H2O, 3 mM K4[FeII(CN)6], 3 mM K3[FeIII(CN)6], 0.5% Triton-X-100, 0.2% Xgal}. Staining solution was removed, and ovaries washed in 0.1% Triton X-100 in PBS for 30 minutes with two changes of buffer. Ovarioles were then mounted in 90% glycerol and imaged on a Zeiss Imager.A2 equipped with an AxioCam MRc, using Zeiss AxioVision software.

To quantify proliferation, dissected ovaries were incubated for 1 hour at room temperature in Grace’s containing 10 μm 5-ethynyl-2′-deoxyuridine (EdU; Invitrogen), immediately preceding fixation and staining. EdU was detected using AlexaFluor-594 via Click-It chemistry, following the manufacturer’s recommendations (Life Technologies). The number of EdU-positive GSCs was measured as a percentage of total GFP-negative GSCs analyzed. Results were subjected to Chi-Square analysis.

Supplementary Material

supplement

Highlights.

  • E78 is required for Drosophila female fertility.

  • Loss of E78 results in decreased niche size and fewer germline stem cells.

  • E78 mutants display increased cyst death during follicle development.

  • E78 cooperates with ecdysone signaling to control cyst viability during oogenesis.

Acknowledgments

Many thanks to Paul Lasko, Henry Krause, Acaimo González-Reyes, the Bloomington and Harvard Exelixis Stock Centers, and the Developmental Studies Hybridoma Bank for fly stocks and antibodies, and the TRiP at Harvard Medical School for pVALIUM vectors and valuable protocols. This work was supported by National Institutes of Health R01 GM069875 (D.D.-B), National Institutes of Health National Research Service Award F32 GM086031 (E.T.A.), and the East Carolina University Division of Research and Graduate Studies and Thomas Harriot College of Arts and Sciences (E.T.A.).

Footnotes

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