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Endocrinology, doi:10.1210/en.2007-1123
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Endocrinology Vol. 149, No. 1 5-14
Copyright © 2008 by The Endocrine Society

Evidence that Androgens Regulate Early Developmental Events, Prior to Sexual Differentiation

Denise R. Goldman-Johnson, David M. de Kretser and John R. Morrison

Centre for Reproduction and Development (D.R.G.-J., D.M.d.K., J.R.M.), Monash Institute of Medical Research, and Australian Research Council Centre of Excellence in Biotechnology and Development (D.M.d.K.), Monash University, Victoria 3168, Australia

Address all correspondence and requests for reprints to: Dr. Denise Goldman-Johnson, The National Institute for Medical Research, The Ridgeway, Mill Hill, London NW7 1AA, United Kingdom. E-mail: djohnso{at}nimr.mrc.ac.uk.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Androgen signaling is critical for normal fetal development but is not thought to regulate events in early embryogenesis. Given the interest in factors controlling the differentiation of embryonic stem (ES) cells, we have explored the possibility that androgens may play a role. This study demonstrates expression of androgen receptor (AR) RNA and protein in four independent mouse ES (mES) cell lines, and shows that the AR is functional and can interact with transfected androgen response elements to promote green fluorescent protein expression. AR mRNA was detected throughout 10-d differentiation in embryoid bodies (EBs). Exposure of EBs to testosterone (T) or dihydrotestosterone, at doses of 1 and 0.1 µM, respectively, promoted formation of beating cardiomyocytes. Flow cytometric analyses demonstrated a significant increase in the number of {alpha}-actinin and tropomyosin (cardiac markers) positive cells after these treatments. Addition of flutamide (1 µM) to T-treated EBs inhibited the T-induced proliferation of cardiomyocytes, confirming that, in this instance, androgens act via the classical AR-mediated genomic pathway. We also report that mES cells express key steroidogenic enzymes, as detected by RT-PCR, and during 24-h incubations secrete T at concentrations of 1.38 ± 0.22 nM, levels comparable to those secreted by cultured Leydig cells. These novel data demonstrate the capacity of androgens to stimulate increased differentiation of mouse ES cells to cardiomyocytes, and are in keeping with recent observations that AR-deficient mice exhibit cardiac impairment in adulthood.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ANDROGENS PLAY AN essential role in the development and differentiation of male genitalia (1), the sexual differentiation of the brain (2, 3), behavioral characteristics (4), and muscle maintenance throughout development. However, to date, there has been little to directly implicate androgens in developmental events prior to the formation of the adrenal glands and testis. Recently, Sauter et al. (5) examined the expression of a range of steroid hormone receptors in mouse embryonic stem (mES) cells. They reported the expression of androgen receptor (AR) mRNA in mES cells, followed by a clear up-regulation of expression on d-1 mES cell differentiation as embryoid bodies (EBs), after leukemia inhibitory factor (LIF) withdrawal. Expression of the AR was maintained throughout 16-d differentiation. Furthermore, Chang et al. (6) have reported AR mRNA expression in preimplantation blastocysts and the inner cell mass, and AR protein expression in mES cells. They also report that AR mRNA expression levels increase with differentiation of mES cells. These early studies have not explored the source of androgens, if any, that explain expression and modulation of the AR at this early developmental stage, nor whether androgen activity promotes differentiation to specific lineages. Here, we further characterize the expression pattern and functionality of the AR at the earliest stages of mouse embryogenesis. In view of the recent observation that mice with a targeted disruption of the AR develop cardiac abnormalities in adulthood (7), we have examined the influence of androgens on the differentiation. We show that androgens enhance differentiation of mES cells to cardiomyocytes and that mES cells are capable of androgen production.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reagents
Cell lines used were: embryonic stem (mES) male w9.5 (8), mES male MPI-II (9), mES male ZIN40 (10), mES female MPI-VI (9), LNCaP (11), and COS7 (12). The hormones testosterone (T), dihydrotestosterone (DHT), and flutamide (F) were obtained from Sigma-Genosys (Castle Hill, Australia) in powder form, and resuspended in filtered AR-grade ethanol to a concentration of 1 mM. Antibodies used were: AR-N20 (Dako, Glostrup, Denmark); OCT3/4 (Santa Cruz Biotechnology, Santa Cruz, CA); SSEA1 (Santa Cruz Biotechnology); biotinylated goat antirabbit (Dako); horseradish peroxidase-conjugated streptavidin (Dako); mouse anti-β-tubulin monoclonal antibody (CHEMICON, Temecula, CA); streptavidin fluorescein isothiocyanate (FITC) (Dako); streptavidin-R Phycoerythrin (PE) (BD PharMingen, San Jose, CA); mouse-anti-Flk1-PE conjugated (BD PharMingen); anti-{alpha}-actinin antibody (Sigma-Aldrich, St. Louis, MO); anti-tropomyosin antibody (Sigma-Aldrich); and goat-antimouse FITC (Dako).

Cell culture
The culture of w9.5, MPI-II, ZIN40, and MPI-VI mES cell lines (13), LNCaP cells (11, 14), and COS7 cells (12), have been previously described. Culture of long-term feeder-free (FF) w9.5 cells and short-term FF MPI-II, ZIN40, and MPI-VI mES cells was as described (13). For all studies, fetal bovine serum (FBS) was charcoal stripped to remove T and performed as previously described (15). Feeder depletion of mES cells before further assaying or growth as FF was performed via trypsinization and re-plating of cells on 0.1% gelatin-coated tissue culture flasks in DMEM with 15% FBS and 1000 U/ml LIF (CHEMICON). After 40-min incubation at 37 C, feeder cells preferentially adhere to the gelatinized flask, and floating mES cells are removed. Cells were re-plated on gelatinized flasks, allowed to expand, and when approximately 80% confluent, the feeder depletion process was repeated. Cells were then either harvested for RNA/protein extraction or re-plated as appropriate for further studies. After the primary feeder depletion step, feeder contamination was found to be 1.7% (±0.33, n = 3) of the purified mES cells as determined by trypan blue exclusion. After the secondary feeder depletion step, feeder contamination was never observed. The viability of cells in all studies was confirmed using trypan blue exclusion. In addition, the effect of all hormones (and relevant ethanol controls) used in the differentiation studies on cell viability after 10-d culture (with passages) was assessed by trypan blue exclusion. This was performed at a range of doses.

EB differentiation assays
Mouse ES cells were allowed to spontaneously differentiate in vitro as previously described (16). For all studies, FBS was charcoal stripped to remove T, performed as previously described (15). Treatments began immediately upon removal of cells from exposure to LIF and were maintained for 10-d differentiation. At d-5 differentiation, individual floating EBs were picked into wells of a gelatin-coated 96-well plate (48 per treatment) and allowed to attach. After 10 d, each well containing an EB was phenotypically scored for the presence of red hemoglobin and contracting cardiomyocytes by personnel without knowledge of the treatments. After optimization, the ideal cell density used in all studies was 2000 cells/ml and a T concentration of 1 µM. The optimized system was used for analysis of the effect of T on the differentiation of w9.5, MPI-II, ZIN40, and MPI-VI mES cells. Each study comprised a control untreated group, ethanol control (1 µl/ml; hormone diluent) group, and treatment group of 1 µM T. In addition, the effects of DHT and the antiandrogen F were analyzed. In differentiation studies, wherever cell dissociation was required (i.e. flow cytometry analysis), trypan blue was used to assess viability. When scoring EBs for beating cardiomyocytes and hemoglobin, dead or dying EBs were excluded from the study.

RT-PCR analyses
Total RNA was extracted from ES cells and EBs using the TRIZOL Reagent (Invitrogen Corp., Carlsbad, CA), DNase treated with the DNA-free kit (Ambion, Austin, TX), and reverse transcribed using Superscript III (Invitrogen Life Technologies, Rockville, MD) according to supplied protocol. PCRs were performed as described (17). All primers were designed to cross exons using Primer3 software (www-genome.wi.mit.edu/cgi-bin/primer/primer3_www.cgi).

AR mRNA was amplified using the primers 3'AR.f (atcccacatcctgctcaag) and 3'AR.r (caaggcactgcagaggagta) to generate a product of 314 bp. To amplify mRNA for six enzymes involved in androgen synthesis the following primers were used: 3'Cyp11A1.f (acttccggtacttgggcttt), 3'Cyp11A1.r(gcttgagaggctggaagttg for 201-bp product; 3'Cyp17A1.f (actgggaagggactgga),3'Cyp17A1.r(ggctagatgtcactgggag) for 315-bp product; 3'3β-HSD.f(tgttggtggaggagaaggatctg),3'3β-HSD.r(tgggtacctttcacattgacgttc) for 253-bp product; 3'17β-HSD.f (atccagagcctcatccattg), 3'17β-HSD.r (aacgccttggaagctgagta) for 164-bp product; 3'5{alpha}-reductase.f (cctggttcctacaggagctg), 3'5{alpha}-reductase.r (cccctgatcagaactggaaa) for 160-bp product; and 3'aromatase.f(agcagcaatcctgaaggaga), 3'aromatase.r(ggaagtactcgagcctgtgc), for 236-bp product. The housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), was detected as a loading control with published primers (18).

Immunochemistry
Western blotting was performed as previously described (19). AR, OCT3/4, and SSEA1 were detected by standard immunohistochemistry as previously reported (20). Antibody concentrations were: AR-N20 (detects mouse, rat, and human AR), 1:20; OCT3/4, 1:50; SSEA1, 1:20; goat-antirabbit biotin-conjugated secondary antibody, 1:500; streptavidin FITC, 1:500; and streptavidin-R PE, 1:25.

Plasmid construction and transfection
The androgen response element (ARE) green fluorescent protein (GFP) vector was constructed using the pEGFP-N1 vector (BD Biosciences Clontech, Palo Alto, CA) backbone. PCR was used to isolate the mouse mammary tumor virus (MMTV)-long terminal repeat from a MMTV-chloramphenicol acetyl transferase vector (21), which comprised the ARE. DNA fragments were ligated into appropriate plasmid vectors, and all derived clones were checked by DNA sequence analysis. For MMTV-long terminal repeat sequence, refer to the GenBank sequence (accession no. X12376). The pEGFP-N1 vector was digested with both EcoRI and AseI to excise the cytomegalovirus promoter, and the plasmid was purified using a Gel Purification kit (QIAGEN, Inc., Valencia, CA). The filtrate was ligated with purified ARE product. The completed vector sequence was checked by DNA sequence analysis. Before transfection, the constructed vector was further purified via cesium chloride ethidium bromide gradient extraction as described (22), and microdialyzed on 0.025 µM filters (Millipore, Billerica, MA) on Tris EDTA for 2 h. The ARE-GFP vector was transiently transfected into feeder depleted w9.5 mES cells using electroporation (22). After overnight recovery, cells were observed for GFP expression after a further 24 h.

Flow cytometry
Flow cytometry was used to assess the number of differentiated cells in each treatment positive for fetal liver kinase-1 (Flk1), {alpha}-actinin, and tropomyosin markers. At d-7 differentiation, floating EBs in control, ethanol control, and 1 µM T-treated groups were analyzed via flow cytometry. Approximately 1 x 106 cells of each treatment group were resuspended in cold flow cytometry buffer [1% BSA wt/vol, 0.01% sodium azide wt/vol in PBS (FLOW)] and stained separately for the three markers. For Flk1 staining, cells were spun down and resuspended in 100 µl cold FLOW buffer with 10% normal mouse serum (Sigma-Aldrich), and incubated with the mouse-anti-Flk1-PE conjugated antibody (1:25) on ice in the dark for 1 h. For both {alpha}-actinin and tropomyosin staining, permeabilization of cells was required. As such, cells were fixed in 2% paraformaldehyde for 20 min on ice and permeabilized in 0.1% Triton-X on ice in the dark for 5–10 min. Cells were then collected and resuspended in cold FLOW buffer with 10% mouse serum and either anti-{alpha}-actinin antibody (1:800) or anti-tropomyosin antibody (1:250) for 30 min on ice in the dark. The cells were resuspended in 0.1% Triton-X with a goat-antimouse FITC secondary antibody (1:100) for 30 min, on ice in the dark, then washed with 0.1% Triton-X, collected, resuspended in 400 µl cold FLOW buffer, and placed on ice in the dark until analysis. Cells were analyzed using a Mo-Flo flow cytometry machine (DakoCytomation, Carpinteria, CA), and results analyzed using Summit software (DakoCytomation) and Prism GraphPad (GraphPad Software Inc., San Diego, CA) for statistics. The readout for the flow cytometry was the percentage of cells sorted as positive for a given fluorescein (or marker).

T assays
FBS (Invitrogen) and Knockout Serum Replacement (KSR) (Invitrogen) were initially assayed for levels of T. Charcoal stripped (CS) media were used to remove T present in the FBS and performed as previously described (15). Media containing CS media were used in the culture of mES cells for T assays (and other studies). The FF w9.5 mES cell line and short-term cultured FF MPI-II, ZIN40, and female MPI-VI mES cells were used to condition CS media for 24 h. Conditioned CS media were sampled after 24 h and frozen for later assaying. T was measured after extraction by RIA using a previously described protocol (23, 24). To ensure that all cells used were ES cells and not differentiated derivatives, after removal of media, cells were karyotyped and immunostained for OCT3/4 and SSEA1.

Briefly, T concentrations in plasma were assessed after organic extraction with a 3:2 mixture of hexane and ethyl acetate. Procedural recoveries were calculated with a parallel sample with tritiated T added. After drying overnight, the organic fraction was reconstituted with 1% gelatin-PBS buffer. The reconstituted aliquots were processed in a RIA with a specific antibody raised to T (Endocrine Sciences T3–125; Endocrine Sciences, Calabasas Hills, CA) and a liquid chromatography purified tritiated T used as a tracer. The cross-reactivity of the T antibody with other steroids was tested by Endocrine Sciences and reported as: DHT (20%), corticosterone (less than 0.01%), estradiol (0.14%), D-1-testosterone (52%), 4-androsten-3b-17b-diol (3%), 5a-androstan-3b-17b-diol (1.8%), D-4-androstenedione (0.5%), and others (less than 0.5%). After 16-h incubation at 4 C, free and bound T were separated with dextran T70-coated charcoal. The T standard was calibrated against a commercial T preparation, and the coefficient of variability within the assay ranged from 3.1–7.5%. The limit of detection for this assay was 0.1 nM.

Statistical analysis
Data were analyzed by one-way ANOVA with a Dunn’s or Tukey’s Honestly Significantly Different post hoc test using Prism 4 software (GraphPad Software). All data are expressed as mean ± SEM.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
ES cells have a functional AR
RT-PCR analysis was undertaken to confirm the expression of AR RNA in three separate male mES lines, w9.5, as standard and FF, MPI-II, and ZIN40, and one female, MPI-VI (Fig. 1AGo), all as FF. The increased level of AR mRNA in standard w9.5 vs. FFw9.5 resulted from minor feeder contamination (of 1.7% from 1xfeeder depletion) and is seen due to the sensitivity of RT-PCR. In addition, there was some slight variation in AR mRNA levels between the different cell lines. This was not observed at protein level. AR protein expression, demonstrated in standard w9.5 and FF w9.5 mES cells by both immunoblotting (Fig. 1BGo) and immunohistochemistry (only standard w9.5 shown, Fig. 1CGo), was also confirmed in MPI-II, ZIN40 and female MPI-VI mES cell lines (data not shown). This was consistent with findings from other laboratories (5, 6).


Figure 1
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  		FIG. 1. Characterization of AR expression in mES cells and EB differentiation. A, PCR analysis of AR mRNA expression in mES cell lines. B, Immunoblot of AR protein expression in mES cells. COS7 cells were used as a negative control and whole testis as a positive control. β-tubulin was used as a loading control. C, Immunohistochemistry of OCT3/4 (intracellular), SSEA1 (cell surface), and AR protein expression in w9.5 mES cells. Phase contrast (PHASE), DAPI nuclear staining, FITC (STAIN), and DAPI-FITC (OVERLAY) are shown. White arrows indicate nuclear localization of the AR signal in some cells; red arrows indicate the dispersed expression of the AR in the cytosol of other cells. The dual sites of expression of the AR in the nucleus and cytoplasm in different cells appeared consistent when the experiments were repeated. D, Transient expression of ARE-GFP vector in w9.5 mES cells in the presence of 10–8 M T. No expression of the ARE-GFP vector was observed in transfected cells in the absence of T (–T). The vector is also depicted. E, The appearance of EBs throughout 10-d differentiation and expression of AR mRNA. All scale bars, 100 µm. +, PCR on cDNA; –, PCR on RT negative controls (no RNA); BC, beating cardiomyocytes; H, hemoglobin.

 
The AR was localized to the nucleus in some cells (Fig. 1CGo, white arrows) and cytoplasm in others (Fig. 1CGo, red arrows). Although the AR is commonly thought to reside in the nucleus, it has also been suggested that it is localized in the cytoplasm and enters the nucleus via a hormone-dependent import process (25, 26). In addition, the cellular localization of the AR may be tissue specific or affected by experimental technique, such as fixation and staining (27). Such contradictory evidence on cellular localization has been reported with other nuclear receptors.

To demonstrate that the androgen pathway was functional in mES cells, a GFP construct driven by an ARE promoter was transfected into w9.5 mES cells, which resulted in robust fluorescence in the presence of androgens (Fig. 1DGo). Thus, mRNA, protein, and gene reporter studies all indicate the presence of a functional AR signaling pathway in mES cells.

Androgens promote the differentiation of mES cells toward cardiomyocytes
To determine the role of the AR in a model of differentiation, AR mRNA expression was monitored at d 3, 5, 7, and 10 in w9.5 EBs (Fig. 1EGo). RT-PCR demonstrated the presence of AR mRNA at all stages, and the signal appeared to intensify as differentiation progressed.

The presence of hemoglobin and beating cells (assumed to be cardiomyocytes) was assessed in the EB system at d 10 of differentiation, in the presence and absence of T. Long-term exposure of ES cells to T below, and including, doses of 10–5 M did not affect cell viability (Fig. 2AGo). This was similarly observed for other hormones and in EBs after 10-d differentiation. A dose-response study identified that the formation of cardiomyocytes was optimal at 1 µM T (Fig. 2BGo), with a cell density of 2000 cells/ml (Fig. 2CGo). Starting cell densities at 3000 and 4000 cells/ml, and similarly at 6000, 7000, and 8000 cells/ml, all resulted in a significant increase in the formation of cardiomyocytes after treatment with 1 µM T (data not shown). Although treatment with 1 µM T at 9,000 and 10,000 cells/ml starting densities always resulted in improved cardiomyocyte yields relative to controls, the large variations in final numbers between repeats did not render this increase statistically significant at n = 3.


Figure 2
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  		FIG. 2. Optimizing culture conditions. A, mES cell viability after 48-h exposure to hormones at various doses. B, Dose-response curve for T on cardiomyocyte derivation. C, The effect of optimization of cell density and 1 µM T on cardiomyocyte formation in EBs. For all studies, n = 3 and *, P < 0.05; **, P < 0.01. Eth, Ethanol.

 
Treatment with the optimal conditions resulted in a significant increase (P < 0.001; n = 14) in the percentage of EBs containing beating cardiomyocytes relative to controls (Fig. 3AGo). The percentage of w9.5 EBs containing beating cardiomyocytes increased from 19.76 ± 2.68 and 20.38 ± 2.95% in control and ethanol control, respectively, to 35.51 ± 4.41% (P < 0.001; n = 14) after T treatment. The percentage of EBs containing hemoglobin was unchanged (Fig. 3BGo). Importantly, these results were observed in the two other male mES cell lines (Fig. 3CGo), with percentages of MPI-II EBs containing beating cardiomyocytes increasing from 25.44 ± 4.03 and 24.32 ± 3.04% in control and ethanol control, respectively, to 53.33 ± 6.3% (P < 0.05; n = 3) after T treatment, and in ZIN40 EBs increasing from 14.67 ± 2.1 and 15.2 ± 0.6% in control and ethanol control, respectively, to 30.57 ± 0.68% (P < 0.05; n = 3) after T treatment. In the female cell line MPI-VI, the same effect of T was observed (Fig. 3CGo) with percentages of EBs containing beating cardiomyocytes increasing from 20.85 ± 4.34 and 18.77 ± 5.25% in control and ethanol control, respectively, to 45.85 ± 1.1% (P < 0.05; n = 3) after T treatment.


Figure 3
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  		FIG. 3. The effect of T treatment on mES cell differentiation. A, Treatment of differentiating w9.5 mES cells with 1 µM T at 2000 cells/ml significantly increases the percentage of EBs with beating cardiomyocytes compared with both control and ethanol control groups (P < 0.001; n = 14). B, Treatment with 1 µM T does not affect the percentage of EBs with hemoglobin. C, Treatment of MPI-II, ZIN40 and female MPI-VI mES cells with 1 µM T significantly increases the percentage of EBs with beating cardiomyocytes compared with both control and ethanol control groups (for all cell lines: *, P < 0.05; n = 3). D, Treatment of w9.5 mES cells with 1 µM T does not affect the numbers of cells positive for the general mesoderm marker Flk1 (Left) but does significantly increase the number of cells positive for {alpha}-actinin (**, P < 0.01; n = 3) and tropomyosin (***, P < 0.001; n = 3), both cardiac cell markers.

 
There was some small variation in the effect of T treatment between cell lines, i.e. the number of EBs with beating cardiomyocytes increased by approximately 57% in w9.5, 47% in MPI-II, 49% in ZIN40, and 43% in the female MPI-VI. These variations may be attributed to minor differences between cell lines and additionally, the number of experimental repeats, i.e. 14 replicates on the main cell line w9.5 and triplicates for all others. Nevertheless, all cell lines showed a statistically significant increase in cardiomyocyte formation with T treatment.

Flow cytometry further confirmed that T promoted the differentiation of cardiomyocytes (Fig. 3DGo). T treatment did not stimulate an increase in the number of cells positive for Flk1, a general marker of mesoderm (n = 3, each group), however, significant increases in the percentage of {alpha}-actinin positive cells were noted (P < 0.01; n = 3). T also increased the percentage of tropomyosin positive cells (P < 0.001; n = 3). Assessment of the duration of T treatment indicated that 10-d exposure to T was necessary to stimulate cardiomyocyte formation (Fig 4AGo), with treatment for the first or second 5-d period resulting in unchanged numbers.


Figure 4
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  		FIG. 4. The effect of time of addition of T, DHT, and F on differentiating w9.5 mES cells. A, The effect of time of addition of 1 µM T on the percentage of EBs with beating cardiomyocytes. EBs were treated for either the first 5 d (d 1–5), last 5 d (d 5–10), or the entire 10 d (d 1–10) of differentiation with 1 µM T. Treatment for the full 10 d was required to induce a significant effect. B, DHT treatment at 0.1 µM significantly increased the percentage of EBs with beating cardiomyocytes relative to all controls, similarly to treatment with 1 µM T (for both *, P < 0.05; n = 3). C, The significant effect (**, P < 0.01; n = 3) of 1 µM T treatment to the percentage of EBs with beating cardiomyocytes is reduced to levels similar to controls when cotreated with 1 µM F (1 µM T +1 µM F). Treatment of EBs with 1 µM F alone reduces the percentage of EBs with beating cardiomyocytes significantly below control and ethanol (Eth) control levels (*, P < 0.05; n = 3). Eth 0.1 µM, ethanol control for 0.1 µM hormone treatment; Eth 1 µM, ethanol control for 1 µM hormone treatment.

 
Androgens act via a classical intracellular AR
The use of DHT, a nonaromatizable androgen, in the EB differentiation assay promoted a significant (P < 0.05; n = 3) stimulation of cardiomyocyte formation at a dose of 0.1 µM DHT (Fig 4BGo). Cotreatment of differentiating cells with T and F, a pure androgen antagonist, at equivalent concentrations (1 µM), demonstrated that F was able to block the androgen effect (Fig. 4CGo) (P < 0.01; n = 3). The treatment of EBs with 1 µM F alone reduced the percentage of EBs containing beating cardiomyocytes significantly below both control levels, to 1.39 ± 1.39% (n = 3), suggesting the possibility that it may be blocking the action of an endogenous source of androgens.

Although the FBS used in all studies was charcoal stripped, the level of T present was assessed directly to exclude the possibility that this source of androgens makes a significant contribution to the studies. T was measured directly in standard FBS, KSR, and CS FBS (Table 1Go). T levels were significantly lower in the FBS relative to the KSR (P < 0.001; n = 9), which was further reduced by charcoal stripping. In an attempt to minimize the impact of media-derived androgens, KSR and standard FBS were not used for any other studies. CS FBS (15%) was used for all studies, resulting in a contribution of subnanomolar levels of T to the media, a level that we believe is unlikely to have made a significant impact on the EB studies.


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  		TABLE 1. T assays of media components

 
Mouse ES cells synthesize and secrete T
The conversion of cholesterol to T requires a coordinated expression of at least four genes (28). RT-PCR demonstrated the expression of the genes encoding steroidogenic enzymes essential to androgen synthesis, namely cytochrome P45011 A1 (Cyp11A1), cytochrome P45017 A1 (Cyp17A1), 3β-hydroxysteroid dehydrogenase (HSD), 17β-HSD, and 5{alpha}-reductase (5{alpha}-Red) (reviewed in Ref. 29), in all mES cell lines (Fig. 5Go). The expression of aromatase mRNA was also demonstrated to show that these cells may potentially be capable of converting T to estrogen.


Figure 5
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  		FIG. 5. Expression of mRNA encoding enzymes essential for androgen synthesis in mES cell lines. GAPDH was used as a loading control. +, PCR on cDNA; –, PCR on RT negative controls (no RNA).

 
T was measured by RIA after separation of mES cell conditioned media by HPLC and demonstrated that at 24 h, FF w9.5 mES cells secreted T at 1.38 ± 0.22 nM, significantly higher concentrations than those found in unconditioned media (P < 0.05; n = 4) (Table 2Go). This finding was further validated in both MPI-II and ZIN40 FF mES cells. Interestingly, although T levels present in media conditioned by female ES cells were higher than in media alone, at 0.53 ± 0.13 vs. 0.3 ± 0.09 nM, respectively, this increase was not significant (Table 2Go).


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  		TABLE 2. T secretion rates from mES cell lines

 
To ensure that all cells used were ES cells and not differentiated derivatives, after removal of media, cells were karyotyped and immunostained for OCT3/4 and SSEA1. All male cells used in the studies were karyotypically normal and positive for both markers of pluripotency (data not shown). Female cells were positive for both markers of pluripotency but of the karyotype 39X0, common in female ES cells (30) (data not shown).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This study demonstrates for the first time that mES cells secrete T, as shown by the measurement of T after HPLC separation. The production of T by male mES cells is also supported by the demonstration of key steroidogenic enzymes in these ES cells. Furthermore, we show the presence of a functional AR in mES cells and determine, for the first time, that androgens, acting through the classical genomic pathway, stimulate the differentiation of mES cells toward a cardiac lineage. As measured by the cardiac-specific markers, {alpha}-actinin and tropomyosin, flow cytometry studies provided unambiguous confirmation that T is affecting an increase in the derivation of cardiomyocytes. The ability of DHT to show a similar degree of stimulation to T (at 10-fold lower concentration) suggests that the action on cardiomyocyte formation is unlikely to be mediated by conversion to estradiol. This conclusion is further supported by the action of F, which blocks the interaction of T/DHT with the AR. We have shown that the AR is clearly present in these cells, confirming earlier reports (5, 6). Although the levels of hormones used to induce the effects on differentiation are above the normal physiological range, e.g. in vivo, T circulates at levels between 6.9 and 39.7 nM in adult men and 0.7 and 2.8 nM in adult women (28), artificial systems such as these often require the concentrations of additives to be superphysiological to stimulate a response (see Ref. 31).

To determine whether the T effect on cardiomyocyte differentiation was resultant from action before or after gastrulation, variations in the time of addition of T were explored. EBs treated for either the first 5 d of differentiation (before gastrulation) or the last 5 d (after gastrulation) did not show significant increases in the formation of beating cardiomyocytes at d 10 (Fig. 4AGo). As such, we conclude that to demonstrate the effect of T on cardiomyocyte formation, treatment must be maintained for the entire period of differentiation.

Our studies also show that mES cells can synthesize T endogenously, at levels comparable to that of cultured Leydig cells (Table 2Go) (32, 33). The capacity of F to suppress cardiomyocyte formation in the absence of added T supports these views. Overall, these results highlight the importance of androgens in early developmental events and, in particular, in the stimulation of cardiomyocyte formation.

The steroid hormone family, other hormones and growth factors, and early embryogenesis
Our data indicate that the novel effects described in this paper are due to specific androgen actions. The interaction between other related hormones and factors and early embryogenesis have been previously reported.

The estrogen receptor RNA is expressed in both human and mouse ES cells (20, 34, 35); treatment with estrogen supports proliferation of ES cells and drives differentiation toward a neuronal lineage (36, 37, 38, 39). Treatment of ES cells with progesterone stimulates growth and proliferation (39), but specific lineage support has not been reported. Furthermore, although the glucocorticoid receptor RNA is expressed in both human and mouse ES cells throughout 16-d mES cell differentiation as EBs (5, 20), dexamethasone drives both human and mouse ES cells toward a hepatocyte lineage (34).

Retinoic acid influences differentiation of ES cells (35), inducing differentiation toward endoderm (31), dorsal interneurons, neuronal progenitors (37, 38, 39), and adipocytes (40), but inhibiting cardiomyocyte formation, in mouse EBs. Furthermore, suppression of the retinoic acid receptor enhances cardiomyocyte gene expression (41, 42, 43).

Classical anabolic hormones include androgens, estrogens, insulin, the IGF family, and GH. Anabolic hormones improve growth and differentiation of cells and increases in body size, a process that involves synthesis of complex molecules.

Insulin is a commonly used supplement in media for both ES cell maintenance and differentiation induction. Supplementation of media with insulin (commonly as ITS-X) results in promotion of cell growth and allows a reduction in concentrations of FBS added to the media (44). Insulin has activated the phosphoinositide 3-kinase pathway, which is important for the maintenance of pluripotency, growth, and survival of mES cells (45). This pathway also activates mammalian target of rapamycin, reported as essential for mES cell and early embryonic proliferation.

It is noteworthy that IGF-I and IGF-II have stimulated cardiomyocyte derivation from human ES cells via the phosphoinositide 3-kinase/Akt signaling pathway (46). Furthermore, the secretion of IGF-I by mES cells injected into Id knockout blastocysts, displaying multiple cardiac defects, rescued them from midgestation lethality (47). Injection of IGF-I and mES cells to a cardiac ischemic region of an adult mouse demonstrably improves cardiac function, whereby IGF-I promotes the differentiation of the mES cells to functional cardiomyocytes (48). Interestingly, androgens are capable of stimulating production of the IGF-I receptor in prostate cancer cell lines, via activation of a nongenomic AR signaling pathway (49). IGF-I also directly activates the AR in the absence of androgens in prostatic tumor cell lines (50). It is possible that androgens and IGF-I interact and work in concert to affect cardiomyocyte differentiation; further investigations would be of interest.

Implications for androgen biology in embryogenesis and cardiac development
A recent study showed that in an AR-deficient mouse model (7), the absence of androgen signaling resulted in a reduced heart-to-body weight ratio, suggesting a developmental defect. This study also reported alterations in ventricular function in response to a challenge with angiotensin II as well as exaggerated cardiac fibrosis. These data, together with our findings, imply that androgens, acting through the AR, play an important role both in cardiac development and in the ability of the cardiac tissue to respond to stress in adulthood (7). What has not been established is whether the susceptibility to disease in AR-deficient mice represents a developmental event or is a reflection of the absence of androgen processing in the adult. Although most authors have tended to focus on androgen function in the adult, our studies suggest that it may be prudent to evaluate androgen action on embryonic events.

Furthermore, the recent study by Beqqali et al. (51) further supports the involvement of androgens in human cardiac development in which, using genome-wide transcriptional profiling, they identified SRD5A2L2 as a novel cardiac-enriched gene from human ES cell differentiation. The novel SRD5A2L2 gene is most similar to steroid 5{alpha}-Red (isoform 2), which encodes the rate-limiting enzyme in the synthesis of DHT from T, and is localized to androgen target tissues such as genital skin and prostate (52, 53). It was reported that the SRD5A2L2 gene was strongly expressed in beating cardiomyocytes, restricted to the heart in d-8.5 post-fertilization mouse embryos, and predominantly in the vena cava and pulmonary vein of a 14-wk-old human fetal heart (51).

Clearly, the androgen pathway is not essential for the anatomical development of the heart because it does develop in humans and mice lacking androgen signaling. However, a high degree of degeneracy often surrounds essential biological processes to ensure viability of the organism. This may be the case for cardiac development and explain why defective androgen signaling does not result in a more pronounced phenotype. In an analogous model, IGF-I similarly appears to play only a supporting role in the formation of cardiomyocytes because the ablation of this gene allows for the normal development of a functional heart (54). Presumably, both androgens and IGF-I are part of a coordinated set of agents that contribute to the optimal development of the heart and its function. What is not clear is whether the absence of one of these factors leads to the formation of a compromised heart that is not competent to deal with the accumulated stresses experienced over the lifetime of an individual.

Mouse ES cells synthesize and secrete T
The demonstration that mES cell lines secrete T challenges our concepts of the source and role of androgens during development. First, our data suggest that mouse ES cells must be added to the short list of cells with the capability of steroid biosynthesis, namely, cells of the ovary, testis, and adrenal cortex. Second, it is clear from our data that androgens have the ability to influence differentiation well before their classical role in sexual differentiation. Androgens may play many other roles in this early embryonic phase. For instance, ES cells have been shown to be immune privileged (55, 56, 57), and perhaps this quality is derived from the synthesis and secretion of androgens, which are known immunosuppressive agents (58). It has been shown that androgens interact with receptors on the plasma membrane of T cells and macrophages to induce a rapid nongenomic response via increases in free Ca2+ fluxes (59, 60). Although this signaling is known to result in immunosuppression, the precise mechanisms of action have not been well defined. Further investigation is required to clarify the contribution of androgens to the apparent immunoprivileged status of mES cells.

Although the early secretion of androgens is highly novel, other studies have shown the secretion of progesterone and 17β-estradiol during human ES cell differentiation (61). From a study by Gerami-Naini et al. (61), it was concluded that EB differentiation is associated with consistent secretion of placental hormones, thus providing evidence that these steroid hormones play an important role in early embryogenesis. Other reports indicate that in vitro fertilized human embryos secrete estradiol and progesterone (62, 63, 64, 65), predominantly when the embryo is cultured with 10–6 M T (65). It would be of interest to assess whether, in this system, EBs are capable of secreting T, particularly because treatment with F alone reduces cardiomyocyte derivation and is presumably inhibiting endogenous androgens. The differentiation of presumptive trophectoderm from ES cells has been reported in mouse and human (61, 66). In view of the capability of the placenta to produce steroids, these placental precursor cells may be the source of hormone secretion in differentiating EBs.

Sex differences in androgen signaling during early embryogenesis
The MPI-VI female mES cells used in these studies display genetic instability, particularly with the rapid loss of one X chromosome. This could have an important effect, not only on the differentiation of these cells, but also on their ability to process androgens as the AR is X linked. It has been reported that the paternally derived X chromosome in females retards development (67), and, as such, if these cells retained an X chromosome from this source alone, their growth rates and differentiation capabilities may be stunted. However, the female mES cell line used here did display pluripotency and normal growth/morphology in culture; thus, it was used in further analyses. In any case, due to the genetic instability of this MPI-VI female mES cell line, it would be necessary to repeat these experiments on other female lines. Ideally, we would use normal XX mES cell lines, but although normal, stable human ES cell lines are readily available, such is not the case with mouse ES cells (68). Where stable XX mES cell have been produced, it has been reported that DNA methylation is globally reduced (69). This phenomenon can be abolished with the elimination of one X chromosome, which, thus, explains the propensity of X-chromosome instability and loss in other female mES cell lines.

When media conditioned by the female mES cell line were assayed for T, levels present were not significantly different from that in control, unconditioned media. This finding differs from that seen with all three male mES cell lines. Although female mES cells express mRNA for genes encoding enzymes essential to androgen formation from cholesterol and additionally, 17β-estradiol formation from T, it is possible that transcription of these genes differs between the two sexes of cell lines. As such, female mES cell lines may not possess the machinery required for androgen synthesis at this stage of embryogenesis. This requires further assessment.

Despite the difference in T secretion, differentiating female mES cells responded to androgen treatment in a manner comparable to that of the male mES cell lines, with significant increases in the derivation of beating cardiomyocytes. This result suggests that the AR in male and female embryos is capable of responding similarly to androgens at this stage of development.

In summary, we have described the unique action of androgens at the earliest stages of embryonic development. We have shown that androgens not only affect in vitro differentiation but are also in fact secreted from ES cells. These findings add a new dimension to the roles played by androgens during development, at stages far earlier than previously thought.


   Acknowledgments
 
We thank Professor D. Handelsman and M. Jimenez (Australian and New Zealand Army Corps Research Institute) for performing all testosterone assays. We also thank Dr. E. Ball (Monash Institute of Medical Research) for donating the mouse mammary tumor virus-chloramphenicol acetyl transferase vector, Dr. A. Voss (The Walter and Eliza Hall Institute of Medical Research) for the MPI-II and MPI-VI mouse embryonic stem cells, Stem Cell Sciences for the ZIN40 cells, and Professor G. Risbridger (Monash Institute of Medical Research) for the LNCaP cells. We thank Dr. E. Stanley (Monash Immunology and Stem Cell Laboratories) for his continued assistance.


   Footnotes
 
This work was supported by National Health and Medical Research Council Australia (Program Grant 334011).

Disclosure Statement: The authors have nothing to disclose.

First Published Online October 4, 2007

Abbreviations: AR, Androgen receptor; ARE, androgen response element; CS, charcoal stripping; Cyp11A1, cytochrome P45011 A1; Cyp17A1, cytochrome P45017 A1; DHT, dihydrotestosterone; EB, embryoid body; ES, embryonic stem; F, flutamide; FBS, fetal bovine serum; FF, feeder-free; FITC, fluorescein isothiocyanate; Flk1, fetal liver kinase-1; FLOW, 1% BSA wt/vol, 0.01% sodium azide wt/vol in PBS; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GFP, green fluorescent protein; HSD, hydroxysteroid dehydrogenase; KSR, Knockout Serum Replacement; LIF, leukemia inhibitory factor; mES, mouse embryonic stem; MMTV, mouse mammary tumor virus; PE, Phycoerythrin; 5{alpha}-Red, 5{alpha}-reductase; T, testosterone.

Received August 13, 2007.

Accepted for publication September 19, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Evidence of a Role for Androgens in Embryonic Stem Cell Function and Differentiation
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