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Identification of a pituitary ERα-activated enhancer triggering the expression of Nr5a1, the earliest gonadotrope lineage-specific transcription factor
Epigenetics & Chromatinvolume 12, Article number: 48 (2019)
Gonadotrope lineage differentiation is a stepwise process taking place during pituitary development. The early step of gonadotrope lineage specification is characterized by the expression of the Nr5a1 transcription factor, a crucial factor for gonadotrope cell fate determination. Abnormalities affecting Nr5a1 expression lead to hypogonadotropic hypogonadism and infertility. Although significant knowledge has been gained on the signaling and transcriptional events controlling gonadotrope differentiation, epigenetic mechanisms regulating Nr5a1 expression during early gonadotrope lineage specification are still poorly understood.
Using ATAC chromatin accessibility analyses on three cell lines recapitulating gradual stages of gonadotrope differentiation and in vivo on developing pituitaries, we demonstrate that a yet undescribed enhancer is transiently recruited during gonadotrope specification. Using CRISPR/Cas9, we show that this enhancer is mandatory for the emergence of Nr5a1 during gonadotrope specification. Furthermore, we identify a highly conserved estrogen-binding element and demonstrate that the enhancer activation is dependent upon estrogen acting through ERα. Lastly, we provide evidence that binding of ERα is crucial for chromatin remodeling of Nr5a1 enhancer and promoter, leading to RNA polymerase recruitment and transcription.
This study identifies the earliest regulatory sequence involved in gonadotrope lineage specification and highlights the key epigenetic role played by ERα in this differentiation process.
Nr5a1 gene (also called Sf-1 or Ad4BP) is a transcription factor (TF) belonging to the nuclear receptor superfamily. In mammals, Nr5a1 is expressed notably in testes, adrenal glands, ventromedial hypothalamic nucleus (VMH) and anterior pituitary gland where it participates in embryonic cell differentiation and adult function . The anterior pituitary is composed of six hormone-secreting cell types, i.e., corticotrope, melanotrope, somatotrope, lactotrope, thyrotrope and gonadotrope cells, originating from common precursor stem cells of the Rathke’s pouch . Several TFs are known to promote pituitary stem cells differentiation into a specific endocrine lineage: POU1F1 is mandatory for the thyrotrope, somatotrope and lactotrope lineages , TBX19 for the corticotrope , PAX7 for the melanotrope  and Nr5a1 for the gonadotrope lineage . During gonadotrope cells specification, Nr5a1 is the earliest specific marker gene known to be expressed  initiating transcription of key genes such as Gnrhr (GnRH receptor gene) and Lhb (β-subunit of the gonadotropin LH gene). As a consequence, mutations in the human Nr5a1 gene  and Nr5a1 knockout in mice lead to gonadotrope deficiency .
Nr5a1 expression depends on tissue-specific cis-regulatory elements. Two main promoters, 1A and 1G, have been characterized. The 1G promoter is the predominantly activated promoter in the pituitary . However, it is not able, alone, to initiate Nr5a1 expression . Additional distal enhancers are required for tissue-specific transcription. Four specific enhancers have been identified that control expression of Nr5a1 in the VMH , fetal adrenal glands , fetal Leydig cells  or gonadotrope cells . The gonadotrope enhancer has been suggested to be implicated in Nr5a1 expression in gonadotropes from mouse embryonic day 13.5 (E13.5) onwards . In a previous work , we characterized the epigenetic marks decorating cis-regulatory regions of Nr5a1. We used a set of three cell lines recapitulating three stages of gonadotrope differentiation. The αT1–1 cells are likely derived from E12.5 common precursor cells from which originate thyrotrope and gonadotrope lineages. These cells do not express Nr5a1 yet. The αT3–1 cells are likely derived from cells engaged in a gonadotrope cell fate at E13.5 and express some of the gonadotrope-specific genes including Nr5a1. The LβT2 cells are likely derived from mature gonadotrope cells and express all the known marker genes [15,16,17]. We observed that although Nr5a1 is already expressed in αT3–1 cells, the epigenetic marks on the gonadotrope enhancer indicate that it is repressed, suggesting that it does not regulate Nr5a1 expression at early steps of gonadotrope specification.
In this work, using functional genomic in vitro and in vivo approaches, we demonstrated that Nr5a1 expression is triggered by another early activated enhancer at the emergence of gonadotrope lineage. We showed that this enhancer is activated by the estrogen pathway through ERα leading to P300 histone acetyltransferase recruitment. We also demonstrated that ERα protects the enhancer from inhibition by DNA methylation and chromatin compaction. Finally, we showed that the enhancer interacts with the Nr5a1 pituitary promoter and increases histone acetylation, RNA polymerase recruitment and Nr5a1 transcription. Activation of this enhancer is thus the earliest known mechanism implicated in gonadotrope cell specification.
Differential chromatin accessibility in Nr5a1 locus during gonadotrope specification
We performed an assay for transposase-accessible chromatin with high-throughput sequencing analysis (ATAC-seq) in αT1–1, αT3–1 and LβT2 cell lines, allowing the identification of new potential cis-regulatory sequences (Fig. 1a and Additional file 1). We observed massive changes in chromatin accessibility in the three cell lines with about 20,000 specific accessible regions per line. Genomic regions associated with genes known to be expressed in the three cell lines, such as Cga or Isl1 promoters, were open in all cell lines. Regions associated with Gnrhr promoter were accessible in both αT3–1 and LβT2, whereas those associated with Lhb promoter were only found in LβT2 cells (Additional file 1). Chromatin accessibility is thus consistent with the maturation stage of these cellular models.
We then analyzed chromatin accessibility of the Nr5a1 locus. In αT1–1, consistent with the absence of Nr5a1 expression, very few genomic regions were accessible in the Nr5a1 locus. In αT3–1, several open chromatin regions could be observed and among them two had already been described: the 1G promoter, in agreement with Nr5a1 expression and, more surprisingly, the fetal Leydig enhancer (FL enhancer). This enhancer has been formerly described to be active specifically in the fetal testis . The gonadotrope enhancer (named hereafter β enhancer) showed only very limited chromatin accessibility in αT3–1 cells, in agreement with our previous observations . The chromatin was also accessible for a previously undescribed region encompassing two very close peaks in intron 4 (peak-1: mm9 chr2:38,559,896–38,560,286 and peak-2: mm9 chr2:38,562,187–38,562,583), named hereafter the α region. This region exhibited only limited chromatin accessibility in αT1–1 cells. In LβT2 cells, the 1G promoter as well as the FL and β enhancers all showed accessible chromatin conformation. In contrast, the chromatin of the α region was no more accessible.
These results strongly suggested that the Nr5a1 locus displays dynamic chromatin accessibility during gonadotrope lineage differentiation. In order to investigate whether the same dynamics could be observed in vivo, we set up an ATAC assay followed by qPCR on developing pituitaries of mouse embryos (Fig. 1b). We observed that the 1G promoter was significantly open at E13.5 and E14.5 compared to E12.5, in agreement with the Nr5a1 expression dynamics during mouse pituitary development . Chromatin at the α region was strongly and transiently accessible at E13.5. While chromatin at the β enhancer displayed a very limited accessibility at E13.5, it was fully open at E14.5. Finally, the FL enhancer was open from E13.5 onwards. Differential chromatin accessibility in Nr5a1 locus during gonadotrope specification was thus validated in developing embryos in vivo.
Discovery of an undescribed early enhancer in Nr5a1 locus
In order to further characterize regions with chromatin accessibility, we studied in the three cell lines the deposition of H3K4me3, H3K4me1 and H3K27ac histone marks, specific for promoters, enhancers and active elements, respectively (Fig. 2a–c). Only the 1G promoter was significantly enriched with H3K4me3 in αT3–1 and LβT2 cells (Fig. 2a). All other regions had significant H3K4me1 enrichment at these three stages of gonadotrope maturation, indicating that they all are potential enhancers (Fig. 2b). Concerning deposition of the active chromatin mark (Fig. 2c), the 1G promoter and FL enhancer were enriched in H3K27ac only in αT3–1 and LβT2 cells. While the β enhancer was decorated with H3K27ac only in LβT2, the α potential enhancer was significantly decorated with H3K27ac in αT1–1 and αT3–1 cells (Fig. 2c, fold enrichment of 2.6 ± 0.4, p < 0.05 and 8.4 ± 1.0, p < 0.01, respectively).
Potential cis-regulatory activity was then studied in luciferase reporter system (Fig. 2d). We observed that the FL enhancer and 1G promoter were active in the three cell lines. A significant cis-regulatory activity of the α enhancer peak-2 could be already observed in αT1–1 that was strongly increased in αT3–1 (fold induction of 5.6 ± 0.8 and 38.5 ± 1.4 over control Pluc–Prl, p < 0.01 in αT1–1 and αT3–1, respectively) but lost in LβT2 cells. As peak-1 did not show any regulatory activity in the αT3–1 cell line (Additional file 2A), subsequent experiments were performed on the peak-2 only and α enhancer will thereafter refer to this peak. The β enhancer displayed a significant cis-regulatory activity only in LβT2 cells (fold induction of 3.8 ± 0.2, p < 0.01). Altogether these results suggest that: (i) the β enhancer is active only during the terminal maturation stage; (ii) the FL enhancer might also be recruited in gonadotropes; (iii) the α region is a genuine gonadotrope enhancer potentially activated in precursor and immature cells.
The α enhancer regulates Nr5a1 expression specifically in immature gonadotropes
CpGs DNA methylation of the active enhancers was investigated by bisulfite DNA sequencing. As shown in Fig. 3a, CpGs of the α enhancer were mostly hypermethylated in progenitors, fully demethylated in immature and methylated again in mature cells. The FL enhancer was hypermethylated in αT1–1 and unmethylated in αT3–1 and LβT2 cells. Thus, both DNA methylation and histone decoration show that the FL enhancer is totally inactive in progenitors and active in immature and mature cells. In contrast, the α enhancer is in a bivalent state in progenitors, exhibiting both active and inactive marks, active in immature and totally silent in mature gonadotropes.
To further confirm the regulatory role of the α enhancer in Nr5a1 expression, the peak-2 genomic sequence was excised from αT3–1 genome by CRISPR/Cas9. Excision was also performed for the FL enhancer. Deletion of the FL enhancer did not alter Nr5a1 expression in αT3–1, whereas deletion of α enhancer sequence led to a strong reduction in expression as revealed by a 85% decrease in Nr5a1 mRNA level (Fig. 3b, p < 0.001). This drastic reduction was observed in three independent clones for two independent gRNA couples, ruling out potential gRNA off target effects. (The genomic sequences of deleted clones along with gRNAs position are shown in Additional file 2B.) In order to inactivate this enhancer without altering the DNA sequence, we targeted dCas9–LSD1, a lysine-specific histone demethylase to the α enhancer. In αT3–1 cells, this led to an 80% decrease in Nr5a1 mRNA level as compared with control gRNA (Fig. 3c, p < 0.01). Decommissioning the α enhancer in LβT2 cells did not impair Nr5a1 expression, while targeting LSD1 to the β enhancer strongly reduced it (fold decrease of about 90%, Additional file 3A). The transient α enhancer activation is thus mandatory for Nr5a1 expression specifically in immature gonadotropes. Chromatin interaction between the α enhancer and 1G promoter was investigated in αT1–1 and αT3–1 using quantitative chromatin conformation capture (3C) assay (Fig. 3d). We also tested potential interactions with control regions, external (Ce) and internal (Ci) to the Nr5a1 locus as well as the FL enhancer. No interaction between the α enhancer and the upstream region (Ce/α) could be detected in the two cell lines. In contrast, in αT3–1 cells (and to a lesser extent in the αT1–1), the α enhancer interacts significantly with the 1G promoter as well as with the FL enhancer (Fig. 3d).
ERα controls cis-regulatory activity of the α enhancer
In order to understand the mechanisms regulating α enhancer activity, genomic sequence conservation of the α enhancer peak-2 was analyzed across mammals (Fig. 4a). A 65-bp core sequence (mm9 chr2:38,560,050–38,560,114) is conserved including a 13-bp stretch showing more than 60% of conservation. According to the cisBP online library , this element corresponds to a perfect ERE motif. No other conserved binding site could be identified in the rest of the core sequence (Additional file 2C).
In order to test the involvement of this 65-bp core sequence in the α enhancer cis-regulatory activity, truncated sequences were tested by luciferase reporter assay in αT3–1 cells maintained in complete steroid-containing medium (Fig. 4b). The full-length α enhancer construct (Pluc–α enh) displayed a significant cis-regulatory activity as compared to the minimal prolactin promoter (Pluc–Prl) used as control. The 65-bp core sequence (Pluc–α enh +65) showed similar activity as Pluc–α enh. Furthermore, deletion of the 65-bp core sequence (Pluc–α enh Δ65) or mutation of the potential ERE motif (Pluc–α enh MutERE) completely abolished cis-regulatory activity in αT3–1 cells. This demonstrates that the 65-bp core conserved sequence is sufficient alone to drive α enhancer activity and that the ERE motif is crucial for this activity.
Both estrogen receptors α (ERα) and β (ERβ) bind to ERE motif. Esr1 and Esr2 mRNAs were quantified by RT-qPCR (Fig. 4c). While Esr2 transcripts were undetectable, Esr1 was expressed in αT1–1 and αT3–1 cells, with a fivefold higher expression level in αT3–1 cells (Fig. 4c, p < 0.01). In mature LβT2 cells, Esr1 mRNA could hardly be detected.
ChIP using an anti-ERα antibody was then performed in αT1–1 and αT3–1 cells. ERα was strongly enriched on the α enhancer as compared to the control region in αT3–1 cells (Fig. 4d, p < 0.001). Binding of ERα to the α enhancer was also observed to a lesser extent in αT1–1 cells (Additional file 4A, p < 0.01).
To further analyze the role of ERα in the regulation of α enhancer activity, Esr1 expression was knocked down using Esr1 SiRNA. Efficiency in ERα decrease was validated by western blot (Additional file 5A). Specific knockdown of ERα abolished α enhancer cis-regulatory activity in αT3–1 cells (Fig. 4e, p < 0.001). A significant decrease in α enhancer activity was also observed in αT1–1 cells (Additional file 4C).
In order to test ligand dependency, α enhancer activity was measured in the presence of 17β-estradiol (E2) or the widely used antagonist ICI 182,780 in the αT3–1 cells. While E2 dose-dependently activated α enhancer cis-regulatory activity, ER inhibition led to a dose-dependent repression (Fig. 4f). A similar repression could be observed using the ERα-specific antagonist, MPP dihydrochloride (Additional file 5B).
To confirm α enhancer dependency to both ERα and E2, activities of the WT- and ERE-mutated α enhancers were assessed in the presence of E2 combined with ERα over-expression in LβT2 cells. We observed that α enhancer activity can be significantly induced by ERα over-expression alone (fold induction compared to basal of 19.7 ± 3.5, p < 0.001, Additional file 3B) and further increased by ERα over-expression combined with E2 treatment (fold induction compared to basal of 65.1 ± 5.4, p < 0.001, Additional file 3B). Mutation of the ERE abolished ERα and E2 effects on α enhancer cis-regulatory activity.
In order to investigate whether ERα is sufficient to activate the endogenous α enhancer in mature gonadotrope cells, ERα was over-expressed in LβT2 cells and the α enhancer chromatin accessibility was assessed by ATAC-qPCR. We observed that over-expression of ERα is sufficient to significantly increase α enhancer chromatin accessibility in mature gonadotropes (fold increase in chromatin accessibility of 2.2 ± 0.4, p < 0.01 Additional file 3C).
These results altogether demonstrate that the α enhancer is active only in immature gonadotrope cells and that this activity is regulated by ERα and E2.
ERα controls Nr5a1 expression through epigenetic regulation of the α enhancer and 1G promoter
To specifically inhibit ERα binding to the α enhancer, the ERE motif of α enhancer genomic sequence was excised using CRISPR/Cas9 in immature gonadotropes. Two αT3–1 clones (ΔERE) bearing homozygous deletion of this ERE were retrieved. As expected, the deletion encompassed the ERE plus four additional bases at each side (Fig. 5a). ChIP assay showed that ERα enrichment on α enhancer was dramatically decreased in ΔERE clones as compared to WT (99% decrease p < 0.001) (Fig. 5b).
Nr5a1 expression level was then quantified in ΔERE clones. Compared to WT clones, Nr5a1 expression was decreased by 85% in ΔERE clones (Fig. 5c, p < 0.001).
To further characterize α enhancer regulation by ERα, epigenetic remodeling at the α enhancer and 1G promoter was then investigated in ΔERE clones. DNase I hypersensitivity assay revealed that excision of ERE-binding site led to a 90% decrease in α enhancer chromatin accessibility (Fig. 5d, p < 0.001). Comparison of histone mark decoration of α enhancer between ΔERE and WT clones revealed that deletion of ERE did not impact H3K4me1 deposition on the α enhancer (Fig. 5e). However, a 75% drop in H3K27ac enrichment could be observed (Fig. 5e, p < 0.001).
Binding of the P300 histone acetyltransferase to the α enhancer was then investigated by ChIP using anti-P300 antibody. P300 was significantly enriched on the α enhancer in αT3–1 and to a lesser extent in αT1–1 cells (Additional file 4B, p < 0.01). Deletion of ERE led to an 80% decrease in P300 recruitment in αT3–1 cells (Fig. 5e, p < 0.001). CpGs DNA methylation was analyzed in ΔERE compared with WT clones (Fig. 5g). Inhibition of ERα binding led to a hypermethylation of the α enhancer in immature gonadotropes.
Analysis of the 1G promoter epigenetics revealed that α enhancer repression did not affect H3K4me3 mark deposition. It, however, decreased H3K27ac enrichment on the 1G promoter (60% decrease p < 0.01, Fig. 5e) without decreasing P300 recruitment. This decrease was associated with a decrease in serine 5-phosphorylated RNA polymerase II recruitment to the TSS (60% decrease p < 0.001, Fig. 5e).
Altogether, these results demonstrate that ERα binding to the α enhancer leads to epigenetic activation of the α enhancer and 1G promoter leading to Nr5a1 expression in immature gonadotropes.
Gonadotrope differentiation is a stepwise process taking place during pituitary development. Gonadotrope lineage is characterized by the expression of Nr5a1, a mandatory TF for gonadotrope cell identity and maturation . However, the molecular mechanisms triggering Nr5a1 expression are still poorly understood. In the current study, we have analyzed the epigenetic mechanisms implicated in this process. Using three cell lines recapitulating different stages of gonadotrope differentiation and combining chromatin accessibility analyses with studies of epigenetic mark deposition and cis-regulatory activity, we re-evaluated the implication of previously described Nr5a1 cis-regulatory sequences and characterized a yet unidentified enhancer element that initiates Nr5a1 expression at the earliest step of gonadotrope specification.
Nr5a1 expression has been suggested to be under the control of the 1G promoter and β enhancer in differentiating gonadotropes [8, 13]. Here, we confirmed that these two regulatory sequences are indeed activated in mature LβT2 cells. However, we showed that although the Nr5a1 1G promoter is already active in immature gonadotropes, this is not true for the β enhancer that does not display any active chromatin marks or cis-regulatory activity at this differentiation stage. This demonstrates that, in immature gonadotropes, molecular mechanisms necessary for β enhancer activity are not yet at work. The β enhancer is thus not the earliest enhancer triggering Nr5a1 expression during gonadotrope specification.
Previous experiments performed by Stallings et al.  demonstrated that genomic fragments encompassing the 1G promoter are not sufficient to induce Nr5a1 expression in vivo in the pituitary. We thus decided to analyze chromatin accessibility in the Nr5a1 locus using our cellular models of gonadotrope differentiation to search for new cis-regulatory elements. Among the different potential cis-regulatory elements showing chromatin accessibility, one enhancer had already been characterized as a regulator of Nr5a1 expression in fetal Leydig cells . We showed that this FL enhancer is active in immature and mature cells and repressed in progenitors. It, however, cannot be implicated in the initiation of Nr5a1 expression as it is located inside the genomic fragments that were tested as negative by Stallings et al. . Interestingly, we identified a previously undescribed enhancer, the α enhancer, with an intriguing activation pattern: It is transiently and specifically activated at the immature stage in vitro. In order to investigate the activation of the α region in vivo, we studied chromatin accessibility in the Nr5a1 locus during mouse pituitary development. Interestingly, the chromatin at the α region was only transiently accessible at E13.5 corresponding to gonadotrope cell emergence and initiation of Nr5a1 expression. Moreover, this α enhancer displays bonafide active enhancer epigenetic mark decorations, with P300 recruitment, H3K27ac deposition and demethylated CpGs. Using both deletion and functional assays, we demonstrated that this enhancer is mandatory for Nr5a1 expression in immature gonadotropes, contrary to the FL and β enhancers. Altogether, these data demonstrate that the α enhancer is the earliest activated cis-regulatory sequence of Nr5a1 gene, and as such, the earliest cis-regulatory sequence specifically activated during gonadotrope specification.
According to our work, Nr5a1 expression would be dynamically regulated by the sequential recruitment of two enhancers, the α enhancer during the early steps of lineage specification and then the β enhancer in mature gonadotropes. The FL enhancer, which is active during both stages, might act as a relay during this process. Supporting this hypothesis, a recent work has demonstrated the involvement of such transiently activated enhancers during the motor neuron differentiation process .
Analysis of the sequence conservation of the α enhancer in mammals allowed us to identify an almost perfectly conserved ERE motif. We demonstrated that ERα binds to this ERE and that regulation of the α enhancer activity is estrogen dependent. During mouse embryogenesis, Esr1 is expressed in the developing pituitary from E12.5 onwards [data from GenePaint.org (image C1253.3.4.B) and Additional file 6A]. However, fetal circulating estrogens are believed to be inactive. Yet, active estrogens can be locally produced either by desulfonation of circulating estrogens sulfate by the steroid sulfatase (STS) enzyme or by aromatization of circulating androgens by aromatase. Genes encoding both enzymes are expressed in mouse and rat adult pituitaries [20,21,22]. Moreover, Sts transcripts (but not of the aromatase coding gene Cyp19a1) are expressed in progenitor and immature gonadotrope cells (Additional file 6B), supporting the idea that differentiating gonadotropes could locally produce the estrogens needed for α enhancer activation.
Interestingly, Nr5a1 expression has been shown to be regulated by estrogens in pituitary and testis [23, 24]. In addition, pituitary Nr5a1 expression can be affected by prenatal exposure to estrogenic endocrine-disrupting chemicals . However, no functional ERE motif was found in Nr5a1 promoters. The α enhancer is thus a bonafide candidate to mediate estrogen regulation of Nr5a1 expression though ERα recruitment.
Here, by precisely deleting ERE motif from the α enhancer genomic sequence using CRISPR/Cas9 strategy, we successfully inhibited ERα binding. We observed that ERα binding is mandatory for Nr5a1 expression in immature gonadotropes. We further demonstrated that ERα binding initiates epigenetic remodeling including P300 recruitment and subsequent H3K27ac deposition, maintaining both an open chromatin state and CpGs hypomethylation. Interestingly, we observed that ERα binding to the α enhancer also remotely regulates the 1G promoter activity. Data obtained using quantitative 3C assay strongly suggest that promoter activation is mediated in part by chromatin looping. Recent evidence obtained from studies in several cellular models [26,27,28] indicates that ERα may regulate gene expression mainly through enhancer activation. It has also been shown in breast cancer cells that ERα binding to enhancers increases P300 recruitment  and decreases DNA CpG methylation . Our data demonstrate that ERα-dependent epigenetic regulation is also crucial during the earliest step of gonadotrope lineage specification.
Interestingly, we observed that ERα and P300 already bind to the α enhancer in progenitor cells and that CpGs around and inside the ERE motif remain hypomethylated. This suggests that ERα binding to the α enhancer in progenitors protects the ERE from de novo CpG methylation and allows enhancer pre-activation by increasing H3K27ac deposition. Very recently, ERα has been shown to be implicated in a biphasic recruitment of P300 on enhancers : Binding of ERα first leads to P300 recruitment and histone acetylation, promoting initial enhancer pre-activation; then, ERα co-activators are recruited to reinforce P300 binding, leading to enhancer maturation and full activation.
Over-expression of ERα combined to E2 treatment in the progenitor cell line is not sufficient to induce Nr5a1 expression (data not shown), suggesting that ERα is not able to activate the endogenous α enhancer in a repressed chromatin environment. This is consistent with several works, showing that ERα requires pioneer TFs to bind to nucleosome-masked ERE sites . Thus, these mandatory pioneer factors might not yet be expressed in the progenitor cells. Based on our study, we can hypothesize that the α enhancer is in a bivalent state in progenitor gonadotropes, silent yet prone to be activated by recruitment of ERα co-activators (Fig. 6). By contrast, over-expression of ERα is sufficient to re-activate the α enhancer in the mature gonadotropes. This suggests that in mature gonadotropes, the absence of ERα expression prevents α enhancer activation although the required co-activators or pioneer factors remain expressed at this stage. Moreover, as ERα is known to be expressed in adult gonadotropes, it would suggest that α enhancer might be recruited again in adulthood to regulate dynamically Nr5a1 expression at important stages of reproductive life.
Molecular mechanisms implicated in α enhancer activation and inhibition are in current investigation and should lead to the identification of new actors of gonadotrope lineage specification and function.
Gonadotrope cell specification is still not well understood, and although key TFs and signaling pathways have been identified, the molecular events implicated in the very early steps of this lineage commitment remain elusive. Deciphering the dynamic of the regulatory enhancer network during gonadotrope specification should significantly improve the understanding of this process. This knowledge is critical for efficient reprogramming of stem cells into mature gonadotropes that would have important therapeutic applications.
The αT1–1, αT3–1 and LβT2 mouse gonadotrope cell lines (generously given by P. Mellon, University of California, La Jolla, CA) were grown in monolayer cultures using high-glucose DMEM supplemented with 10% fetal bovine serum (FBS) and 0.2% penicillin/streptomycin, at 37 °C with 5% CO2.
Antibodies were purchased from Abcam: anti-H3K4me1 ab8895; anti-H3K4me3 ab8580; anti-H3K27ac ab4729; anti-PolII S-5-P ab5131; anti-ERα ab32063; anti-KAT3B ab19541, and from Santa Cruz: anti-GAPDH sc-25778.
Assay for transposase-accessible chromatin with high-throughput sequencing analysis was performed as described . Briefly, 50,000 nuclei from αT1–1, αT3–1 and LβT2 cell lines were transposed using Illumina Nextera Transposase. Library fragments were amplified using NEBnext PCR master mix and custom Nextera PCR primers. Sequencing was performed on a NextSeq 500 system at the ICM iGenSeq core facility from Paris. Three independent replicates were done for each line. Data analysis was performed according to Buenrostro et al. . Briefly, ATAC-seq reads were mapped on mouse genome mm9 using Bowtie 2  and peak calling was done using MACS2 . Peaks were then tested for consistency among three independent replicates, and data visualization was done using IVG software .
Pregnant mice of the SWISS background at 12.5, 13.5 and 14.5 days post coitum were purchased from Janvier Labs. Mice were killed by cerebral dislocation and embryos retrieved and anesthetized in ice-cold PBS. Developing pituitaries were dissected under magnification glasses. Six pituitaries from age-matched embryos were pooled and cells dispersed in high-glucose DMEM with 10% FBS supplemented with 1 mg/mL collagenase D and 40 U/mL DNase I for 30 min at 37 °C. Transposition, amplification and libraries were performed as described . The experiment was performed three times independently in triplicates.
For each genomic region, specific enrichments were quantified by real-time PCR using LightCycler 480 Instrument (Roche Diagnostics) and Takyon No ROX SYBR master mix (Eurogentec). Specific primers are described in Additional file 7. Raw qPCR data were normalized to control region and to E12.5 stage of development. The control region, already used in , is located at mm9 chr11:111,296,111–111,296,224 and is a region that displays neither enrichment for histone modifications, TF binding or chromatin accessibility so far on every tested tissues or cell types according to the ENCODE data. Change in chromatin accessibility between E12.5 and E13.5 or E12.5 and E14.5 embryonic stages was compared using a Mann–Whitney test.
DNase I hypersensitivity assay
DNase I-sensitive assay was performed as described . Briefly, 50,000 nuclei were digested with 1 U RQ1 DNase I (Promega) for 5 min at 32 °C. DNA was then extracted by proteinase K treatment and phenol/chloroform extraction. DNA fragments were segregated by size by centrifugation for 24 h at 25,000 g on a 9% sucrose cushion. A 500-µL fraction representing fragments of less than 1000 bp was collected at the top of the gradient and precipitated. The experiment was performed in triplicates on two independent clones. The α enhancer sequence enrichment in the purified DNA fractions was quantified by real-time PCR. Primers are described in Additional file 7.
ChIP experiments were performed as described . Briefly, 20 million cells were cross-linked with 1% formaldehyde for 10 min (or 30 min for P300 ChIP) at 37 °C, and then, formaldehyde was quenched by adding glycine (125 mM final). After nuclei extraction and lysis, chromatin was sheared by five 25-s rounds of sonication at 50% setting with a Bioblock Scientific Vibra-Cell sonicator. About 50 µg of chromatin for histone chromatin epigenetic marks and 100 µg for TFs along with 5 µg of antibodies per immunoprecipitation were combined. Immunoprecipitation was carried out at 4 °C overnight using Dynabeads™ Protein G (Invitrogen, ThermoFisher Scientific). After extensive washings and elution, chromatin cross-linking was reverted by heat and DNA purified using phenol/chloroform extraction and precipitation. Each ChIP experiment was performed at least six times independently.
For each genomic region, specific enrichments were quantified by real-time PCR. Specific primers are described in Additional file 7. Raw qPCR data were normalized to chromatin inputs and control region and were then compared using ANOVA followed by Dunnett’s multiple comparison, performed independently for each cell line and each histone marks or TF after checking for normal distribution using Kolmogorov–Smirnov test.
Luciferase reporter assay
Cloning of the cis-regulatory region: Genomic regions encompassing each potential cis-regulatory element were amplified from mouse DNA and cloned upstream from a minimal prolactin promoter (Prl) in the pGL3-basic vector (Promega) as previously described . Truncated regions were obtained by PCR or fusion PCR. Primers for amplifications and truncations are described in Additional file 7.
Cell transfection: Briefly, 50,000 cells were transiently transfected in 96-well plates with 100 ng/well of pGL3 plasmids along with 5 ng/well of pRL-SV Renilla plasmid as an internal control for normalization, using Lipofectamine® 2000 (ThermoFisher Scientific) according to the manufacturer’s protocol. When indicated, cells were co-transfected with either Dharmacon ON-TARGETplus non-targeting control SiRNAs (D-001810-01-05) or ON-TARGETplus Esr1 SiRNA (LQ-058688-01) at 10 pmol/well. When indicated, 24 h after transfection, cells were treated with either vehicle, E2, ICI 182,780 or MPP dihydrochloride at the indicated concentrations in complete steroid-containing or steroid-deprived medium. At 48 h after transfection, firefly and Renilla luciferase activities were measured using the Dual-Luciferase Reporter Assay System (Promega) according to the manufacturer’s instructions. Each experiment was performed six times independently in quadruplicates. Data were normalized to Renilla and control Pluc–Prl plasmid level for each condition. Normal distribution was checked using Kolmogorov–Smirnov test, and ANOVA followed by Dunnett’s multiple comparison tests was performed independently for each cell line and each condition.
Bisulfite conversion of genomic DNA and sequencing
DNA methylation was performed as described previously . Bisulfited fragments were cloned into pGEM-T Easy vector (Promega). At least five clones per cell line were selected and sequenced to determine the state of CpG methylation. Primers used for bisulfited DNA amplification are listed in Additional file 7.
Quantitative chromatin conformation capture
3C experiments were conducted as described . Briefly, 10 million cells were cross-linked with 2% formaldehyde for 10 min at 25 °C, and formaldehyde was then quenched by adding glycine (125 mM final). After nuclei extraction, restriction using 750 U of HindIII was carried out overnight at 37 °C under gentle agitation. After enzyme inactivation, in nucleus ligations were performed overnight at 16 °C. Chromatin cross-linking was reverted by heat and DNA purified using phenol/chloroform extraction and precipitation. A BAC library containing all possible chimeric fragments in equal amount was generated as followed: About 10 µg of RP23 225F7 Nr5a1 BAC (gift from Dr CT. Gross, EMBL, Italy) was restricted with HindIII, religated and used as a control PCR template. For each cell line, four independent experiments were performed in triplicates. Chimeric DNA fragments quantification was carried out using specific primers encompassing the HindIII restriction sites on both the α enhancer and tested regions by qPCR. The BAC library was used as a standard for unbiased concentration estimation. Data were normalized to the α enhancer and internal control Ci chimeric fragment level and analyzed using Kolmogorov–Smirnov test and ANOVA followed by Dunnett’s multiple comparison tests.
CRISPR/Cas9 deletions and dCas9–LSD1 enhancer inhibition
pLV hUbC-Cas9-T2A-GFP (#53190), pLV hUbC-dCas9-T2A-GFP (#53191) and pSPgRNA (#47108) plasmids  were purchased from Addgene. The Lsd1 coding sequence was amplified from pET15B-hLSD1 (gift from Dr. Y. Shi, Harvard Medical School, Boston, USA) and cloned in frame of the C terminus of dCas9 in the pLV hUbC-dCas9-T2A-GFP plasmid. The specific guide RNAs were designed using CCTop online tool  and cloned in pSPgRNA plasmid.
About 5 million αT3–1 cells were electroporated with the Cas9 expression vector and sgRNA plasmids at a ratio of 1.5:8.5 µg using a Neon® Transfection System (Invitrogen) according to the manufacturer’s protocol (two pulses at 1500 mV for 15 ms). The empty pSPgRNA was used as control. At 48 h post-transfection, GFP-positive cells were sorted using the FACS Aria II on the PIC2 facility of the Unit of Functional and Adaptive Biology.
For enhancer deletion studies, sorted cells were plated at low density and expanded. Single colonies were tested for deletion by PCR. A minimum of two independent homozygous clones were selected.
For enhancer decommissioning assay, sorted cells were directly processed for RNA extractions. These experiments were performed in triplicates, three times independently.
RNA extraction and mRNA quantification
Total RNA was isolated from cells using Trizol reagent according to the manufacturer’s protocol. RNAs (1 µg) were reverse transcribed with SuperScript II reverse transcriptase (Invitrogen) using random primers according to the manufacturer’s instructions. Specific primers for qPCR quantification are described in Additional file 7. For each cell line, experiments were conducted four times independently in triplicates.
Protein extraction and western blot
Cellular proteins were extracted in Laemmli buffer and separated on a 10% SDS–PAGE. After transfer, nitrocellulose membranes were incubated with anti-ERα (1:1000) or anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH; 1:3000) antibody in Tris-buffered saline containing 0.01% Tween 20 (TBS-T) supplemented with 5% milk overnight at 4 °C. After extensive washing, blots were incubated with a horseradish peroxidase-conjugated secondary antibody (GE healthcare #NA934V) in TBS-T/5% milk for 60 min at a room temperature and then washed. Immunodetection was performed using an enhanced chemiluminescence detection system (GE Healthcare).
Immunohistochemistry staining of ERα
E12.5, E13.5 and E14.5 mouse embryos were proceeded for paraffin-embedded classical histology and were sectioned into 5-µm thickness. The sections were mounted on positively charged slides. The slides were deparaffinized using Histolemon and re-hydrated. Epitopes retrieval was performed by incubation in 0.05% citraconic anhydride buffer (pH 7.4) at 100 °C for 15 min. Endogenous peroxidase was inhibited by incubation in 30% H2O2 for 20 min. Endogenous biotin molecules were blocked with endogenous avidin/biotin blocking kit (ab64212), and non-specific binding was blocked by incubation with 10% goat serum diluted in PBS for 1 h. Subsequently, the sections were incubated overnight in a humid chamber at RT 1 h with ERα antibody (1/200; ab32063 abcam). Sections were extensively washed and incubated for 1 h with the biotinylated anti-rabbit IgG secondary antibody (1/500; ab97049 abcam). Sections were extensively washed and incubated 1 h with HRP-conjugated streptavidin (1/1000; ab7403 abcam). The sections were finally treated with diaminobenzidine in the dark, washed, then rapidly counterstained with Mayer’s Hemalun and mounted in Eukitt medium.
Availability of data and material
The datasets used and analyzed during the current study are available from the corresponding author on reasonable request.
Schimmer BP, White PC. Minireview: steroidogenic factor 1: its roles in differentiation, development, and disease. Mol Endocrinol. 2010;24(7):1322–37.
Rizzoti K, Lovell-Badge R. Early development of the pituitary gland: induction and shaping of Rathke’s pouch. Rev Endocr Metab Disord. 2005;6(3):161–72.
Simmons DM, Voss JW, Ingraham HA, Holloway JM, Broide RS, Rosenfeld MG, et al. Pituitary cell phenotypes involve cell-specific Pit-1 mRNA translation and synergistic interactions with other classes of transcription factors. Genes Dev. 1990;4(5):695–711.
Pulichino A-M, Vallette-Kasic S, Tsai JP-Y, Couture C, Gauthier Y, Drouin J. Tpit determines alternate fates during pituitary cell differentiation. Genes Dev. 2003;17(6):738–47.
Mayran A, Khetchoumian K, Hariri F, Pastinen T, Gauthier Y, Balsalobre A, et al. Pioneer factor Pax7 deploys a stable enhancer repertoire for specification of cell fate. Nat Genet. 2018;50(2):259–69.
Ingraham HA, Lala DS, Ikeda Y, Luo X, Shen WH, Nachtigal MW, et al. The nuclear receptor steroidogenic factor 1 acts at multiple levels of the reproductive axis. Genes Dev. 1994;8(19):2302–12.
Lin L, Philibert P, Ferraz-de-Souza B, Kelberman D, Homfray T, Albanese A, et al. Heterozygous missense mutations in steroidogenic factor 1 (SF1/Ad4BP, NR5A1) are associated with 46, XY disorders of sex development with normal adrenal function. J Clin Endocrinol Metab. 2007;92(3):991–9.
Kimura R, Yoshii H, Nomura M, Kotomura N, Mukai T, Ishihara S, et al. Identification of novel first exons in Ad4BP/SF-1 (NR5A1) gene and their tissue- and species-specific usage. Biochem Biophys Res Commun. 2000;278(1):63–71.
Stallings NR, Hanley NA, Majdic G, Zhao L, Bakke M, Parker KL. Development of a transgenic green fluorescent protein lineage marker for steroidogenic factor 1. Endocr Res. 2002;28(4):497–504.
Shima Y, Zubair M, Ishihara S, Shinohara Y, Oka S, Kimura S, et al. Ventromedial hypothalamic nucleus-specific enhancer of Ad4BP/SF-1 gene. Mol Endocrinol. 2005;19(11):2812–23.
Zubair M, Ishihara S, Oka S, Okumura K, Morohashi K. Two-step regulation of Ad4BP/SF-1 gene transcription during fetal adrenal development: initiation by a Hox-Pbx1-Prep1 complex and maintenance via autoregulation by Ad4BP/SF-1. Mol Cell Biol. 2006;26(11):4111–21.
Shima Y, Miyabayashi K, Baba T, Otake H, Katsura Y, Oka S, et al. Identification of an enhancer in the Ad4BP/SF-1 gene specific for fetal Leydig cells. Endocrinology. 2012;153(1):417–25.
Shima Y, Zubair M, Komatsu T, Oka S, Yokoyama C, Tachibana T, et al. Pituitary homeobox 2 regulates adrenal4 binding protein/steroidogenic factor-1 gene transcription in the pituitary gonadotrope through interaction with the intronic enhancer. Mol Endocrinol. 2008;22(7):1633–46.
Laverrière J-N, L’Hôte D, Tabouy L, Schang A-L, Quérat B, Cohen-Tannoudji J. Epigenetic regulation of alternative promoters and enhancers in progenitor, immature, and mature gonadotrope cell lines. Mol Cell Endocrinol. 2016;15(434):250–65.
Alarid ET, Windle JJ, Whyte DB, Mellon PL. Immortalization of pituitary cells at discrete stages of development by directed oncogenesis in transgenic mice. Development. 1996;122(10):3319–29.
Windle JJ, Weiner RI, Mellon PL. Cell lines of the pituitary gonadotrope lineage derived by targeted oncogenesis in transgenic mice. Mol Endocrinol. 1990;4(4):597–603.
Xie H, Hoffmann HM, Meadows JD, Mayo SL, Trang C, Leming SS, et al. Homeodomain proteins SIX3 and SIX6 regulate gonadotrope-specific genes during pituitary development. Mol Endocrinol. 2015;29(6):842–55.
Weirauch MT, Yang A, Albu M, Cote AG, Montenegro-Montero A, Drewe P, et al. Determination and inference of eukaryotic transcription factor sequence specificity. Cell. 2014;158(6):1431–43.
Rhee HS, Closser M, Guo Y, Bashkirova EV, Tan GC, Gifford DK, et al. Expression of terminal effector genes in mammalian neurons is maintained by a dynamic relay of transient enhancers. Neuron. 2016;92(6):1252–65.
Milewich L, Garcia RL, Gerrity LW. Steroid sulfatase and 17β-hydroxysteroid oxidoreductase activities in mouse tissues. J Steroid Biochem. 1984;21(5):529–38.
Connolly PB, Resko JA. Estrone sulfatase activity in rat brain and pituitary: effects of gonadectomy and the estrous cycle. J Steroid Biochem. 1989;33(5):1013–8.
Carretero J, Vázquez G, Blanco E, Rubio M, Santos M, Martín-Clavijo A, et al. Immunohistochemical evidence of the presence of aromatase P450 in the rat hypophysis. Cell Tissue Res. 1999;295(3):419–23.
Zheng W, Jimenez-Linan M, Rubin BS, Halvorson LM. Anterior pituitary gene expression with reproductive aging in the female rat. Biol Reprod. 2007;76(6):1091–102.
Majdic G, Sharpe RM, Saunders PT. Maternal oestrogen/xenoestrogen exposure alters expression of steroidogenic factor-1 (SF-1/Ad4BP) in the fetal rat testis. Mol Cell Endocrinol. 1997;127(1):91–8.
Brannick KE, Craig ZR, Himes AD, Peretz JR, Wang W, Flaws JA, et al. Prenatal exposure to low doses of bisphenol A increases pituitary proliferation and gonadotroph number in female mice offspring at birth. Biol Reprod. 2012;87(4):82.
Miranda-Carboni GA, Guemes M, Bailey S, Anaya E, Corselli M, Peault B, et al. GATA4 regulates estrogen receptor-α-mediated osteoblast transcription. Mol Endocrinol. 2011;25(7):1126–36.
Jefferson WN, Kinyamu HK, Wang T, Miranda AX, Padilla-Banks E, Suen AA, et al. Widespread enhancer activation via ERα mediates estrogen response in vivo during uterine development. Nucleic Acids Res. 2018;46(11):5487–503.
Carroll JS, Meyer CA, Song J, Li W, Geistlinger TR, Eeckhoute J, et al. Genome-wide analysis of estrogen receptor binding sites. Nat Genet. 2006;38(11):1289–97.
Li W, Hu Y, Oh S, Ma Q, Merkurjev D, Song X, et al. Condensin I and II complexes license full estrogen receptor α-dependent enhancer activation. Mol Cell. 2015;59(2):188–202.
Fleischer T, Tekpli X, Mathelier A, Wang S, Nebdal D, Dhakal HP, et al. Oslo Breast Cancer Research Consortium (OSBREAC), Børresen-Dale AL, et al. DNA methylation at enhancers identifies distinct breast cancer lineages. Nat Commun. 2017;8(1):1379.
Murakami S, Nagari A, Kraus WL. Dynamic assembly and activation of estrogen receptor α enhancers through coregulator switching. Genes Dev. 2017;31(15):1535–48.
Manavathi B, Samanthapudi VSK, Gajulapalli VNR. Estrogen receptor coregulators and pioneer factors: the orchestrators of mammary gland cell fate and development. Front Cell Dev Biol. 2014;2:34.
Buenrostro JD, Giresi PG, Zaba LC, Chang HY, Greenleaf WJ. Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods. 2013;10(12):1213–8.
Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Methods. 2012;9(4):357–9.
Zhang Y, Liu T, Meyer CA, Eeckhoute J, Johnson DS, Bernstein BE, et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 2008;9(9):R137.
Thorvaldsdóttir H, Robinson JT, Mesirov JP. Integrative genomics viewer (IGV): high-performance genomics data visualization and exploration. Brief Bioinform. 2013;14(2):178–92.
Pincas H, Amoyel K, Counis R, Laverrière JN. Proximal cis-acting elements, including steroidogenic factor 1, mediate the efficiency of a distal enhancer in the promoter of the rat gonadotropin-releasing hormone receptor gene. Mol Endocrinol. 2001;15(2):319–37.
Hagège H, Klous P, Braem C, Splinter E, Dekker J, Cathala G, et al. Quantitative analysis of chromosome conformation capture assays (3C-qPCR). Nat Protoc. 2007;2(7):1722–33.
Kabadi AM, Ousterout DG, Hilton IB, Gersbach CA. Multiplex CRISPR/Cas9-based genome engineering from a single lentiviral vector. Nucleic Acids Res. 2014;42(19):e147.
Stemmer M, Thumberger T, del Sol Keyer M, Wittbrodt J, Mateo JL. CCTop: an intuitive, flexible and reliable CRISPR/Cas9 target prediction tool. PLoS One. 2015;10(4):e0124633.
This work benefited from equipment and services from the iGenSeq core facility, at ICM and the PIC2 facility from the Unit of Functional and Adaptive Biology. We are thankful to Blandine Gausserès for her help with the FACS experiments. We also thank Dr Pamela Mellon who developed and provided us the cell lines, Dr Shi Yang and Madeline Schuck for the gift of pET15B-hLSD1 plasmid and Dr Cornelius Gross and Dr Angelo Raggioli for the gift of the RP23-225F7 BAC. We are grateful to Dr Michel Cohen-Tannoudji for his constructive feedbacks.
This work was supported by grants from the Paris Diderot Paris 7 University, from the Centre National de la Recherche Scientifique (CNRS) and from the Institut National de la Santé et de la Recherche Médicale (INSERM). DL is recipient of a grant from Societé Francaise d’Endocrinologie. VP is recipient of a fellowship from Paris Diderot University.
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