Plasmids and antibodies.

The mouse SF-1 expression plasmids T7-Myc-SF-1 and pCEP4SF-1 were provided by David Moore. The full-length cDNA of mouse LRH-1 was obtained by reverse transcription-PCR from undifferentiated ES cell total RNA. The sequence of PCR primers used are 5′CAAGAATTTCCGCTAAGAATGTCTGAG 3′ (the forward primer) and 5′AGTAACTTCCAGGGGTGC 3′ (the reverse primer). For mammalian expression and in vitro translation, plasmids pHA-mLRH-1 and pGBK-mLRH-1 were generated by insertion of mLRH-1 cDNA into pCMV-HA and pGBK-T7 (Clontech, Sparks, Md.), respectively. TheOct4 luciferase reporter vectors have been described previously (19). TheOct4 reporter PP-Luc contains a 0.4-kb XbaI-BanI fragment from the proximal-promoter region, and PP* is identical to PP except for the point mutations inserted in the DR0 element (19). The reporter PE-PP-Luc contains a 1.2-kb BamHI-BanI fragment from theOct4 promoter region, and PE-PP* is identical to PE-PP except for the insertion of point mutations in the DR0 element (19). PE-SV40-Luc was constructed by the insertion of a 0.8-kb BamHI-XbaI PE fragment of theOct4 enhancer into pGL3-SV40-Luc (Promega, Madison, Wis.). Anti-mSF-1 antibody was purchased from Upstate (Charlottesville, Va.; catalog no. 13702). Anti-Oct4 (no. 8628) and anti-actin (no. sc-1616) antibodies and all of the horseradish peroxidase-conjugated secondary antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, Calif.). The anti-mouse LRH-1 antibody used in Western blot analysis and supershift assays, which was produced in our laboratory, was raised specifically against the N terminus of the recombinant LRH-1 mouse protein expressed in bacteria. The portion of theLRH-1 cDNA encoding the N-terminal 100 amino acids was inserted into a His-tagged bacterial expression vector, pRSET-A (Invitrogen, Carlsbad, Calif.).

Cell culture and transient transfection.

P19 cells and COS-1 cells were maintained in Dulbecco's medium supplemented with 10% fetal calf serum. Wild-type andLRH-1^−/− ES cells were maintained on 0.1% gelatin-coated plates in Dulbecco's medium supplemented with 15% fetal calf serum, 110 μM 2-mercapoethanol, 1,000 U of recombinant murine leukemia inhibitory factor (LIF; Chemicon, Temecula, Calif.), 100 U per ml, of penicillin per ml, and 100 μg of streptomycin per ml. For reverse transcription-PCR using differentiated P19 and ES cells, cells were differentiated with 1 μM RA in the absence of LIF and harvested at the indicated times. For Northern blot analysis using differentiated ES cells, wild-type orLRH-1^−/− ES cells were cultured on 1% gelatin-coated bacterial plates in the absence of LIF. Cells were harvested on the indicated days, and RNA was isolated. Day 0 cells were harvested 24 h after seeding. All the transient transfections were performed with Fugene 6 as specified by the manufacturer (Roche, Indianapolis, Ind.). The expression vector and reporter DNA were cotransfected into P19 cells plated in six-well plates (2 × 10⁵ cells per well) for 24 h prior to transfection. TheRenilla RL-Luc reporter (Promega) DNA was also cotransfected as an internal control to correct for differences in transfection efficiency. Total plasmid DNAs were balanced with empty vector pCMV-HA. After 48 h of incubation, the cells were harvested and the luciferase activity was analyzed using the dual-luciferase assay kit (Promega) as specified by the manufacturer.

Electrophoretic mobility shift assays.

In vitro- translated mLRH-1 and mSF-1 were produced using a T7/TNT in vitro translation kit as specified by the manufacturer (Promega). COS-1-overexpressed proteins and P19 and ES cell nuclear extracts were extracted with 2× binding buffer (25 mM HEPES [pH 7.9], 150 mM KCl, 0.4 mM EDTA, 2 mM dithiothreitol, 20% glycerol, 1 mM phenylmethylsulfonyl fluoride, 1× proteinase inhibitor cocktail [Roche]). The sequence of the DR0 probe and electrophoretic mobility shift assay were performed by to methods described previously (19). The sequence of PE1 was as follows: sense, 5′GCATCCTGGCCATTCAAGGGTTGAGTACTTGTT 3′; antisense, 5′GCTAAACAAGTACTCAACCCTTGAATGGGCCAGG 3′ (−928 to −919). The sequence of PE2 was as follows: sense, 5′CTAGGATTGTCCAAGCCAAGGCCATTGTCCTGCCC 3′; antisense, 5′CTGAGGGCAGGACAATGGCCTTGGCTTGGACAATC 3′ (−867 to −858).

Chromatin immunoprecipitation (ChIP) assay.

Chromatin immunoprecipitation ChIP assays were performed by the on-line protocol provided by the company Upstate. Undifferentiated and differentiated P19 and ES cells were treated with 1% formaldehyde. Cross-linked DNA/protein was extracted and disrupted by sonication. The recovered DNA was amplified with gene-specific primers. The sequences of the primers surrounding the DR0 site in theOct4 PP (from −127 to +95) are CCTCCGTCTGGAAGACACAGGCAGATAGCG for forward and CGAAGTCTGAAGCCAGGTGTCCAGCCATGG for reverse; the sequence of the primers surrounding the PE1 and PE2 sites of the PE of theOct4 gene (from −1000 to −824) are GCTGGGGAAGTCTTGTGTGA (forward) and R:GCTTCCAGCCTAGTTCCTGG (reverse).

Northern blot analysis.

Total RNAs from different time points of differentiated P19 and ES cells were isolated using Trizole reagent (Invitrogen). Northern blot analysis was performed as specified in protocol PT1190-1 (Clontech). Blots were hybridized with³²P-labeled cDNAs corresponding to LRH-1 (bases 712 to 1296 [GenBankNM_030676]), Sox2 (bases 1239 to 1903, [GenBankU31967]), Oct4 (bases 908 to 1260 [GenBankNM_013633]), FGF4 (bases 1 to 609 [GenBankNM_010202]), UTF1 (bases 324 to 918 [GenBankNM_009482]), and REX1 (bases 461 to 2073 [GenBankNM_009556]). To ensure equal loading of RNA samples, the blots were probed with radiolabeled cDNA corresponding to the rat 18S rRNA (18S rRNA, bases 293 to 970 [GenBankX01117]).

Immunofluorescent and immunohistochemical staining.

Collection of blastocysts and immunofluorescent staining were performed by methods described previously (24,28). Female mice were superovulated by treatment with pregnant mare serum gonadotropin and human chorionic gonadotropin, and the blastocysts were flushed out of the uterus at E3.5 to 4.0. The zona pellucida of the embryos was removed by brief exposure to Tyrode's solution (pH 2.1 to 2.5) (Irvine Scientific, Santa Ana, Calif.). The embryos were fixed with 4% paraformaldehyde and permeabilized by incubation with 0.2% Triton X-100 in phosphate-buffered saline (PBS). After being blocked in 10% donkey serum plus 2% bovine serum albumin, the embryos were incubated with normal rabbit and goat immunoglobulin G (IgG) or rabbit anti-LRH-1 antibody and goat anti-Oct4 antibody in blocking buffer at 4°C overnight. They were then washed with PBS and incubated with fluorescein isothiocyanate FITC-conjugated donkey anti-rabbit IgG and Texas Red-conjugated donkey anti-goat IgG (Jackson ImmunoResearch Laboratories, Inc., West Grove, Pa.) and washed with PBS. They were stained with 0.5 μg of Hoechst 33258 (Sigma, St. Louis, Mo.) per ml and mounted on glass slides with Vectashield mounting medium for fluorescence (Vector Laboratories, Burlingame, Calif.) or kept in PBS buffer and examined with an Axioskop 2 immunofluorescence microscope (Carl Zeiss).

The embryos at E6.5 were fixed in 10% formaldehyde in PBS and embedded in parafilm. Antigens were unmasked by boiling the sections (7.5 μm) and treating them with 3% H₂O₂ and were blocked in 10% normal goat serum. An immunostaining kit from Santa Cruz Biotechnology was used to detect the LRH-1 or Oct4 protein. After being stained, the slides were counterstained in 0.05% methyl green to visualize histological structures.

Whole-mount and section in situ hybridization.

Whole-mount in situ hybridization was carried out as described previously (19). TheLRH-1 cDNA fragment corresponding to nucleotides 1155 to 1540 andOct4 cDNA (19) were labeled with digoxigenin (Dig) (In vitro translation kit; Promega) as cRNA probes. E6.5 embryos were fixed in 4% paraformadehyde and embedded in paraffin. Sections were probed with³⁵S-labeled sense or antisense cRNA probes against the LRH-1 cDNA fragment.

Genotyping and identification ofLRH-1^−/− embryos.

TheLRH-1^+/− mice were purchased from Lexicon Genetics Inc. The accession number corresponding to the OmniBank mice that were obtained isNM_030676. Genotypes of weaned mice or embryos were determined by PCR analysis of tail-cut DNA samples or yolk sacs. The primer set for the wild-typeLRH-1 allele (forward F1, 5′ TCTGCTAGTTTGGATACTGG 3′; forward F2, 5′ AAAGGACTGCCAATAATTTCGCT 3′; reverse R1, 5′ TTACAGAGTGAAGTTCCAGG 3′; reverse R2, 5′ AAGTGGATCTCTGAGTCTGAG 3′) was derived from the sequence of the first exon and intron; the reverse primer for the mutantLRH-1 allele was the same as that for the wild-type, and the forward primer (R4, 5′ GCAGCGCATCGCCTTCTATC 3′) was derived from theneo gene within the targeting vector. Genomic DNA was extracted from tails of pups or from the yolk sacs of embryos after E7.0, from the whole blastocysts after immunofluorescent staining, or from scraped embryonic tissues from stained sections. Primers F1 plus R1 and R4 were used to genotype the pups, and the nested PCR strategy was used for genotyping the blastocysts and scraped embryonic tissues.

Generation ofLRH-1^−/− ES cells.

LRH-1^−/− ES cells were generated by deletion of exon 6 on both chromosomes, using a two-step targeting strategy (see Fig.7A). A mouse ES cell line was first generated in whichloxP sites were placed in the introns surrounding exon 6. ES cells that underwent homologous recombination were selected with G418 and then transfected with a plasmid encoding Cre recombinase; clones in which exon 6 and the neomycin resistance gene were excised to allow selection for a second targeting event were identified by Southern blot analysis. These cells were then subjected to a second round of mutagenesis, using a targeting vector in which exon 6 was replaced by the neomycin resistance gene, and were reselected with G418. Southern blot analysis identified clones no longer containing a wild-typeLRH-1 gene. The absence ofLRH-1 transcript was verified by Northern blot analysis (see Fig.7C).

Detection and display of LRH-1/SF-1 response elements in the mouse genome.

A set of 19 LRH-1/SF-1 responses elements (LRE) was collected from the published data (see Table1). These sites were used to train a hidden Markov model (HMM) using HMMER v.2.32 by Eddy (14). All mouse chromosomes were scanned using the HMM with an expected value cutoff set to 0.01, and the chromosomal coordinates of the resulting hits were saved. The chromosomal locations of genes of interest were taken from the KnownGenes tables of the UCSC Genome Browser database (26). The LREs were visualized by uploading the list of hits to the UCSC Genome Browser (27), using the “custom tracks” feature (see Fig.2A).

Expression ofLRH-1 in ES cells.

To investigate whether LRH-1 regulatesOct4 expression during early embryonic development, we comparedLRH-1 andSF-1 expression patterns in ES cells and P19 cells by Northern analysis (Fig.1). As previously shown,SF-1 was coexpressed withOct4 in undifferentiated P19 cells and expression of both genes decreased during differentiation (19).LRH-1 was not expressed in P19 cells. In contrast,LRH-1 andOct4 were coexpressed in undifferentiated ES cells whereasSF-1 was not (Fig.1). Expression ofLRH-1 decreased to undetectable levels in ES cells after 12 h of RA treatment. The expression ofOct4 also decreased with RA treatment and was almost undetectable after 36 h of RA treatment. Thus, Northern blot analysis showed thatSF-1 is selectively expressed in P19 cells andLRH-1 is expressed in ES cells (Fig.1). The concomitant expression pattern ofLRH-1 andOct4 in ES cells suggested that LRH-1 might regulateOct4 expression.

Direct binding of LRH-1 to the Oct4 PE and PP.

It was reported that there are two SF-1 response elements in theOct4 promoter: DR0 (AGGTCAAGGCTA [from −42 to −31]) and the SF1β site in theOct4 PP region (5,19). Expression ofOct4 at the epiblast stage is regulated by the proximal enhancer in addition to the proximal promoter (31,56). Therefore, we searched theOct4 promoter for additional SF1/LRH-1 response elements. To do this, we generated a matrix of known LRH-1-binding sites (Table1) from which a Hidden Markov Model was generated using HMMER and used to search the entire mouse genome. This search identified six LRH-1 binding sites. Site 1 is in a nonconserved region outside the proximal enhancer. Two sites, 2 and 3, are located in the evolutionally conserved region (CR2) that corresponds to theOct4 PE. Sites 4 and 5 (site 5 corresponds to SF1β site in reference5) are in a moderately conserved region, and site 6 is the DR0 element in a highly conserved region (CR1) of the (PP) (Fig.2A).

To address the function of LRH-1 in the regulation ofOct4 expression, we tested the ability of LRH-1 protein to bind to these predicted response elements. We focused on the LRH-1-binding sites in the evolutionarily conserved region, sites 2, 3 and 6, which were termed PE1, PE2, and DR0, respectively. The binding of SF-1 and LRH-1 to the PE1, PE2, and DR0 sites was confirmed by supershift analysis (Fig.2B). Both SF-1 and LRH-1 bound with high affinity to the DR0 and PE2 probes and weakly to the PE1 probe. The anti-SF-1 antibody disrupted the binding of COS1-overexpressed SF-1 proteins with the probes (lanes 2, 6, and 10). The anti-LRH-1 antibody supershifted the binding complex generated by LRH-1 protein (lanes 4, 8, and 12). To detect endogenous LRH-1 DNA-binding activity, we extracted nuclear proteins from undifferentiated ES cells or cells that had been treated with RA for 36 h, and electrophoretic mobility shift assay was performed. A fast-migrating complex appeared when nuclear extracts from undifferentiated ES cells were used; this complex was lost when RA-differentiated cell extracts were used (Fig.2C). LRH-1 antibodies supershifted the complex in ES cells (lane 5), but SF-1 antibodies did not (lane 4). The antibody supershift results agreed with theLRH-1 expression results for ES cells using Northern analysis (Fig.1). As previously reported (19,20), a transient retinoid acid induced factor complex formed with theOct4 DR0 probe in extracts from differentiated P19 cells; a complex with similar mobility was detected using extracts prepared from differentiated ES cells (Fig.2C, lane 2).

To demonstrate binding of endogenous LRH-1 to theOct4 promoter in ES cells, ChIP was employed. The results of ChIP analysis demonstrated direct binding of LRH-1 to theOct4 PE and PP in vivo (Fig.2D). Using Oct4 PP- and PE-specific ChIP primers, strong PCR-amplified signals were generated from undifferentiated ES cell DNA coimmunoprecipitated with anti- LRH-1 antibodies, demonstrating that LRH-1 binds to these two regions of theOct4 promoter in vivo. Similar binding activities were observed in P19 cells with anti-SF-1 antibodies but not anti-LRH-1 antibodies. Binding of LRH-1 and SF-1 decreased on RA-induced differentiation in ES cells and P19 cells, respectively. This DNA-binding pattern is consistent with the expression profiles of SF-1 and LRH-1 in P19 and ES cells (Fig.1). These results clearly demonstrate that endogenous LRH-1 in undifferentiated ES cells is directly bound to theOct4 PP and PE in vivo and support a role for LRH-1 in the regulation ofOct4 expression in ES cells.

LRH-1 activation ofOct4 through the PE and PP.

To determine whether LRH-1 can regulate expression of theOct4 gene, a set of luciferase reporters was constructed as illustrated in Fig.3A and described in Materials and Methods. The various Oct4 luciferase reporter constructs were transfected into undifferentiated and differentiated P19 cells. Insertion of theOct4 PE upstream of the PP enhanced reporter activity about fivefold in undifferentiated P19 cells (Fig.3B). When the LRH-1-binding site in the PP was mutated (PP*), the PE enhancer activity was lost in P19 cells (Fig.3B). Regulation by the Oct4 PE could be bestowed on a heterologous promoter since a similar effect was seen when the PE was placed before the simian virus 40 (SV40) basic promoter (Fig.3B). In RA-differentiated P19 cells, the PE had no enhancer activity in either the wild-type context of the Oct4 promoter or linked to the SV40 promoter (Fig.3B). These results show that transcription factors present in undifferentiated P19 cells but not in differentiated P19 cells (i.e., SF-1) act on theOct4 PE.

To further address the function of SF-1 and LRH-1 on the PE enhancer, we cotransfected SF-1 and LRH-1 expression vectors with the differentOct4 luciferase reporters in RA-treated P19 cells (Fig.3C). The transfection results showed that SF-1 and LRH-1 increased the PP and PE-PP reporter activities fourfold whereas SF-1 and LRH-1 had no effect on the mutated PP* and PE-PP* reporter activities. Regulation of a heterologous reporter by SF-1 and LRH-1 could be bestowed by introduction of the PE into the SV40 promoter. Transfected SF-1 and LRH-1 had no activity on the control SV40 promoter but produced twofold activation of the PE-SV40 reporter. These in vitro results demonstrated that endogenous SF-1 and transfected SF-1 or LRH-1 in P19 cells can activate reporter activity through the proximal enhancer of theOct4 gene by binding to elements in the PE; such activation is dependent on the proximal promoter.

LRH-1 expression during early murine embryonic development.

To investigate the physiological relevance of LRH-1 to the regulation ofOct4 expression in vivo, we examined the expression pattern ofLRH-1 in mouse embryos at early developmental stages when theOct4 gene is expressed. Using LRH-1-specific antibodies generated against the N-terminal 100 amino acids of mouse LRH-1, the inner cell mass (ICM) and primitive endoderm of the blastocyst (E3.5 to E4.0) were specifically immunostained; the trophectoderm layer and blastocoelic cavity were only weakly stained (Fig.4A, panel g). No staining was seen in experiments performed with control preimmune IgG (panel c). At this stage,Oct4 is also strongly expressed in the ICM and primitive endoderm (32) (panel h). The expression pattern of LRH-1 protein in the blastocyst was identical to that of β-galactosidase staining in a previously reportedLRH-1:lacZ knockout mouse model (39). The expression ofLRH-1 in embryos from E6.5 to E7.5 was detected by whole-mount in situ hybridization with Dig-labeledLRH-1 cRNA probe. At the advanced egg cylinder stage (E6.5 to E7.0), LRH-1 mRNA was detected throughout the embryonic and extraembryonic regions, including the visceral endoderm and embryonic ectoderm (Fig.3B, panel a). During the early primitive streak stage embryo (E7.5), LRH-1 mRNA was still expressed throughout the embryo (Fig.4B, panel b). There are conflicting reports ofLRH-1 expression in the embryonic ectoderm (39,45). Therefore, to confirm thatLRH-1 was expressed in the epiblast and to demonstrate coexpression ofLRH-1 andOct4 proteins in the embryonic ectoderm, embryos at E6.5 to E7.0 were dissected and hybridized with cRNA probe againstLRH-1 cDNA and immunostained with anti-LRH-1 and anti-Oct4 antibodies. Results from section in situ showed that LRH-1 was expressed throughout the entire embryo at E6.5 including the embryonic and extraembryonic cell layers (Fig.4C). Consistent with the in situ results, LRH-1 protein was detected throughout the embryo, including the visceral endoderm, embryonic ectoderm, and the extraembryonic region (Fig.4D, panel b). Oct4 was specifically located in the ectoderm of the epiblast (panel c). Thus, theLRH-1 gene is coexpressed with theOct4 gene in the ICM and epiblast during early developmental stages.

Disruption ofLRH-1 gene expression leads to embryonic death.

The in vitro and in vivo results suggested that LRH-1 probably plays a key role in regulatingOct4 expression during early embryonic development. To further understand the regulation ofOct4 expression by LRH-1 in vivo, theLRH-1 gene was disrupted by gene targeting. Disruption of theLRH-1 gene was undertaken by insertion of the β-geo cassette into the first exon ofLRH-1 gene via homologous recombination in ES cells (Fig.5A) (54). Southern blot and PCR analysis demonstrated the insertion of the β-geo gene (Fig.5B and C). A cDNA probe that spanned exons 6 to 9, encoding the ligand-binding domain, was used as probe to detect LRH-1 mRNA in mutant embryos.LRH-1 mRNA was present in wild-type E6.5 to E7.0 embryos but not inLRH-1^−/− embryos, demonstrating that theLRH-1 gene was successfully disrupted (Fig.5D) through the replacement of the first exon with the β-geo gene. The genotype of the embryos was further confirmed by genomic PCR analysis.

Breeding results showed that matings of heterozygous offspring failed to produce pups homozygous for the mutantLRH-1 allele. In total, 248 pups were genotyped (Table2); 119 of the mice were wild type and 129 were heterozygous, but no homozygous pups were detected. These data confirm previous data that disruption of the LRH-1 gene causes embryonic lethality (13,15,39). The lower ratio (1.1:1) ofLRH-1^+/− to wild-type pups (expected 2:1 based on Mendelian genetics) indicates haploinsufficiency (13). To determine precisely when homozygousLRH-1 mutants die, timed-mated heterozygous females were sacrificed at different gestational stages and the blastocysts or embryos were genotyped by PCR analysis (Table2).LRH-1^−/− blastocysts were found at E3.5 to E4.0 at a wild-type/heterozygote/homozygote ratio of 23:42:21, which is close to the expected Mendelian ratio of 1:2:1. TheLRH-1^−/− blastocysts appeared morphologically normal (Fig.6A, panel i). Mutant embryos were also found at E6.5 to E7.0, but the ratio (12%) was much lower than expected. NoLRH-1^−/− embryos were detected at E7.5 to E9.5. These data demonstrate thatLRH-1^−/− embryos die around E6.5.

Loss ofOct4 expression in the LRH-1^−/− embryos.

The activation ofOct4 reporters by LRH-1 and the early embryonic death of theLRH-1^−/− embryos led us to examineOct4 expression in theLRH-1^−/− embryos. Analysis ofOct4 expression showed a strong signal in the ICM of blastocysts at E3.5 to E4.0 and epiblast of wild-type embryos at E6.5 (Fig.4 and6).Oct4 was expressed in the ICM (Fig.6A, panels h and l) in theLRH-1^+/− orLRH-1^−/− blastocysts even though the LRH-1 staining signal was weak (panel g) or absent (panel k), respectively. However, at E6.5 to E7.0,Oct4 mRNA was undetectable in epiblasts ofLRH-1^−/− embryos (Fig.6B, panels c and d). The mutant embryos were much smaller than the wild-type controls. Further analysis revealed that histologic abnormalities inLRH-1^−/− embryos were apparent as early as E6.5 to E7.0 (Fig.6C). At this stage, the extraembryonic and embryonic tissues in wild-type embryos were clearly distinguished from each other (Fig.6C, panels a to c); in contrast, they were difficult to distinguish in theLRH-1^−/− embryos (panels d to f). Moreover, inLRH-1^−/− embryos, the visceral endoderm was expanded and the cells were loosely arranged; the embryonic ectoderm showed a marked disorganization, and the proamniotic cavity was malformed. The loss ofOct4 expression in the epiblast ofLRH-1^−/− embryos was further confirmed by immunohistochemical staining (panel f). These results indicated that LRH-1 is not required to maintain the expression of theOct4 gene in the ICM but is required to maintain its expression in the epiblast of developing embryos.

Rapid down-regulation ofOct4 expression in differentiatingLRH-1^−/− ES cells.

To determine if the effects of loss of LRH-1 onOct4 expression were cell autonomous, anLRH-1^−/− ES cell line was established. Due to the inability to generateLRH-1^−/− ES cell lines from blastocyst outgrowths from our LRH-1 knockout mice, a sequential knockout strategy using two different targeting vectors was used in which exon 6, which encodes the T-box region of the DNA-binding domain and part of the hinge region, was disrupted in bothLRH-1 alleles (Fig.7A). Southern blot analysis confirmed deletions in bothLRH-1 alleles (Fig.7B). As expected, Northern blot analysis of theLRH-1^−/− ES cells failed to detectLRH-1 mRNA in these cells (Fig.7C). Monolayer cultures ofLRH-1^+/+ andLRH-1^−/− ES cells were induced to differentiate for 4, 8, and 12 days by withdrawal of LIF. No RA was added to the ES cell cultures, so that the kinetics of Oct4 repression was slower (comparing Fig.1A with 6C). No differences were seen in the number or morphology of wild-type and knockout cells during differentiation (data not shown).

Expression ofOct4 was compared in wild-type andLRH-1^−/− ES cells induced to differentiate for 4, 8, and 12 days. Oct4 expression decreased much more rapidly inLRH-1^−/− ES cells than in wild-type cells (Fig.6C).LRH-1 expression was lost and the expression ofOct4 decreased more rapidly in the mutant ES cells than in differentiating wild-type ES cells. Expression ofSox2,FGF4,UTF1, andREX1, which are dependent on Oct4 (1,7,33,52), were maintained in wild-type ES cells but dropped inLRH-1^−/− ES cells after removal of LIF (Fig.6C). Thus, in the absence of LRH-1, there is a failure to maintain Oct4 expression with a concomitant loss of expression of genes involved in maintaining pluripotence in ES cells.

Complementary expression and function of SF-1 and LRH-1 in P19 and ES cells.

Based on work with P19 cells, it was previously hypothesized that the orphan receptor SF-1 regulatesOct4 expression. However, the survival ofSF-1 knockout mice till birth demonstrates that SF-1 is not a key factor in the maintenance ofOct4 expression during early embryonic development (30). Expression ofSF-1 in P19 cells appears to be a quirk of their derivation. ES and P19 cells are derived from different embryonic developmental stages: the former is a nontumor cell line derived from blastocyst ICM (about E4.5), whereas the latter is a teratocarcinoma cell line derived from transplanted epiblast stage cells at E7.5 (10,53). P19 cells are considered a representative cell system to study events that occur during gastrulation stages of embryonic development. Unlike P19 cells, ES cells do not express SF-1 but do express LRH-1 (Fig.1 and2C and D), which more faithfully reflects the in vivo situation in mouse embryos (10). Thus, LRH-1 and not SF-1 regulates Oct4 expression during early embryonic development.

Regulation ofOct4 expression.

Despite its crucial role in development and maintenance of the germ cell lineage, relatively little is known about how the Oct4 gene is regulated. Thecis regulation ofOct4 expression throughout the pluripotent life cycle has been defined using a LacZ transgenic reporter system (56). Three important regulatory regions were defined in the Oct4 promoter by this study: the PP element and the PE and DE. The transgenic analysis established that the DE is essential for the expression of Oct4 in all stages of the pluripotent life cycle except the epiblast stage, when Oct4 expression is regulated by the proximal enhancer. The proximal promoter is essential for Oct4 expression at all stages of the pluripotent life cycle. There are likely to be several important transcription factors binding to each of these elements at different stages. To date the Sp factors Sp1 and Sp3 and several nuclear receptors, including retinoid receptors and the orphan receptors COUP-TF and SF-1, have been implicated in the regulation of Oct4 expression (5,6,31,42,51). However, KO mouse models have not substantiated a functional link between any of these factors and regulation of Oct4 expression. In contrast, the orphan receptor GCNF binds to the DR0 element in the PP and regulates Oct4 expression. The functional consequence of inactivation of the GCNF gene is loss of proper repression of the Oct4 gene in somatic cells at gastrulation (19).

In this study we established that the orphan nuclear receptor LRH-1 also regulatesOct4 expression by binding to elements in the PP and PE. The consequence of inactivation of theLRH-1 gene is loss ofOct4 expression at the epiblast stage of development. The requirement of LRH-1 to maintainOct4 expression in ES cells of the epiblast correlates directly with the requirement for the elements to which it binds for epiblast expression (56). Phylogenetic comparison of the sequence of theOct4 promoter shows that the elements regulatingOct4 expression are conserved across species (38). Importantly, the conservation of the DR0, PE1, and PE2 sequences between mice and humans suggests that LRH-1 probably will play an essential role in regulating humanOct4 expression in differentiating ES cells and embryonic development.

Developmental roles ofLRH-1.

The broad expression ofLRH-1 during embryonic development suggests that it probably likely impact multiple developmental processes (17). Belanger's group generated anLRH-1:lacZ knock in reporter allele and observed β-galactosidase staining in heterozygous morulae and blastocysts (39). We have confirmed the expression of LRH-1 protein in the ICM of blastocysts. It was reported thatLRH-1 mRNA was detectable only in the visceral endoderm and not in the embryonic ectoderm from E5.5 to E6.5 (39). In our experiments,LRH-1 mRNA and protein are clearly detectable throughout the entire embryo at E6.5 (Fig.4). This discrepancy may be due to the use of different detection systems. Unfortunately, we were unable to detect the expression of thelacZ reporter gene in our β-geo knock-in allele. This is due to deletion of 70 bp upstream of theLRH-1 ATG initiation codon caused by the insertion of the β-geo gene, which disruptsLRH-1 and LacZ reporter expression.

Inactivation of theLRH-1 gene results in embryonic lethality at a later stage than inOct4 mutant embryos, which die on day E5.5 lacking an ICM as the morula differentiated along the trophectodermal lineage (32). This suggests that LRH-1 is not required to maintainOct4 expression in the ICM of blastocysts but, rather, is required to maintain its expression in the epiblast. Consistent with the results obtained in the embryo,Oct4 expression is maintained in undifferentiatedLRH-1^−/− ES cells. LRH-1 may not play a significant role in early preimplantation development. Its expression in the zygote and morula may simply represent expression of the factor before the time when it is required to maintainOct4 expression.

TheLRH-1 expression pattern at later embryonic stages supports a role for LRH-1 in regulating endodermal development and expression of endodermal genes, such as α-fetoprotein,HNF1α,HNF3β, andHNF4α during early hepatic and pancreatic development (2,29,39,40,49). Likewise, LRH-1 may regulate these genes in the visceral endoderm. It was previously reported that inactivation of theLRH-1 gene leads to defective visceral endoderm development (39). Thus, there are likely to be multiple defects in theLRH-1^−/− embryos that contribute to embryonic lethality.

Role ofLRH-1 in ES cells.

Several pluripotency factors in addition to Oct4, includingSox2,FGF4,UTF1, andREX1, are more rapidly down-regulated upon withdrawal of LIF in theLRH-1^−/− ES cell line than in wild-type ES cells. Previous studies showed that Oct4 functions alone or as a complex with other factors to regulate the expression of these genes (1,7,33,52). Northern analysis of the wild-type ES cells showed that althoughOct4 expression was decreased along withLRH-1 levels after withdrawal of LIF, expression ofSox2,FGF4,UTF1, andREX1 was maintained. These data suggest that there is adequate Oct4 to maintain their expression. However, inLRH-1^−/− ES cells, a more rapid loss ofOct4 expression coincided with decreased expression ofSox2,FGF4,UTF1, andREX1. The decreased expression of these genes is unlikely to be a direct consequence of the absence of LRH-1, since no LRH-1-binding sites have been identified in their promoters. Decreased expression ofSox2,FGF4,UTF1, andREX1 is not a generalized, nonspecific effect due to unhealthy ES cells, since most genes are expressed normally as measured by microarray analysis (data not shown). The early stages of ES cell differentiation probably approximate what is occurring in the epiblast in that pluripotence must be maintained even in the presence of nascent differentiation signals.

Maintenance of Oct4 expression is unlikely to be the only role of LRH-1 in regulating ES cell function. In addition to pluripotence, another property of ES cells is a high proliferative capacity. LRH-1 plays an important role in regulating proliferation through regulation of cyclin G1 expression (9). Interestingly it was found that LRH-1 and β-catenin synergistically regulate intestinal cell proliferation through cyclin G1 (9). Inactivation of the β-catenin gene is lethal to the embryo at the same stage asLRH-1 and presents a similar phenotype (25). Thus, LRH-1 and β-catenin may cooperate to regulate ES cell proliferation and expansion from an ICM to an epiblast.

Novel regulatory switch for Oct4 expression.

Based on our findings in this study and our previous studies, we propose a novel regulatory switch forOct4 gene expression during ES cell differentiation and early embryonic development (Fig.8). LRH-1 activates or maintainsOct4 gene expression in the epiblast of mouse embryos and undifferentiated ES cells. Maintenance ofOct4 expression by LRH-1 is mediated through binding to LRH-1 elements in the proximal enhancer and promoter. Upon differentiation of ES cells or gastrulation of embryos, GCNF expression is induced and LRH-1 is down-regulated, leading to the replacement of LRH-1 by GCNF at theOct4 promoter. This switch in transcription factors blocks the DR0 in the proximal promoter and repressesOct4 gene expression in somatic cells. In the ICM and primordial germ cells, other transcription factors (X) bind to the distal enhancer and proximal promoter to maintainOct4 expression. The reciprocal regulation ofOct4 by GCNF and LRH-1 through binding to common sequence elements has profound implications for the maintenance of pluripotence and differentiation of ES cells. Both LRH-1 and GCNF are orphan nuclear receptors that may be regulated by ligands, which provides another potential mechanism for regulating pluripotency in ES cells. The demonstration that LRH-1 is required to maintainOct4 expression in the epiblast of mouse embryos opens a new window to view ES cell maintenance, pluripotence, and differentiation and a new target with which to manipulate these processes.

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