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Department of Reproduction and Development (A.U., M.P.O., A.G., A.O.B.), and Department of Neurology and Erasmus Center for Biomics (T.M.L.), Erasmus MC, Rotterdam 300 DR, The Netherlands
Address all correspondence and requests for reprints to: Arzu Umar, Erasmus MC, Department of Reproduction and Development, P.O. Box 1738, 3000 DR Rotterdam, The Netherlands. E-mail: a.umar{at}erasmusmc.nl.
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Abstract |
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) was found in response to androgen. Androgen-induced posttranslational modification of RhoGDI1 and glycolytic enzymes may be an important functional link between signaling pathways and cytoskeletal rearrangements in control of growth and development of the male internal genital tract. |
Introduction |
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reduced form of T, is involved in development of the prostate and external genitalia from the urogenital sinus and the tubercle (4, 5). Both T and DHT perform their actions via one and the same intracellular receptor, the androgen receptor (AR), but DHT is a more potent androgen. Binding of either T or DHT to the AR might be a mechanism to regulate distinct androgenic effects in target tissues (5). In humans, the differential action of T and DHT is illustrated in clinical syndromes such as androgen insensitivity (AIS) and 5-reductase type2 deficiency. Complete AIS patients have a nonfunctional AR, which results in the absence of Wolffian duct and urogenital sinus derived structures, and feminized external genitalia (1). In contrast, patients with a 5-reductase type2 deficiency have normally developed epididymides and vasa deferentia but have feminized external genitalia (6). Surprisingly, studies in mice deficient for steroid 5-reductase 1 and 2, have shown that treatment with nonreducible T analogs was sufficient for secondary sex organ formation and growth (7). Therefore, it seems that in rodents, DHT acts largely as a signal amplifier (5). In experimental setups, the synthetic androgen R1881 (methyltrienolone) is often used because it is not further metabolized and exerts both T and DHT effects. The AR is a member of the steroid/nuclear receptor superfamily, and functions as a transcription factor upon androgen binding (8). Structurally, the AR protein can be divided into separate domains, e.g. the N-terminal transcription activation domain, the DNA binding domain, and the C-terminal ligand binding domain. The ligand binding domain contains a strictly ligand-dependent transcription activation function and interacts with nuclear cofactors (9, 10). Upon ligand binding, the AR binds to specific genomic androgen response elements, thereby regulating transcription of specific genes.
AR expression in the developing male genital tract occurs in a strict temporal, and cranial to caudal fashion, which is first detected in mesenchymal cells and later in epithelial cells. Mesenchymal AR is expressed along the rat genital tract as early as embryonic d 14 (E14), whereas epithelial expression in the epididymis starts at E18 in the caput epididymis and after birth in the cauda epididymis and vas deferens (11). Thus, initiation of androgen-dependent differentiation of the Wolffian duct system into epididymis and vas deferens occurs before epithelial cells express a detectable level of AR protein. During development, mesenchymal cells are important androgen targets that elicit androgenic effects in epithelial cells via paracrine factors and mesenchymal-epithelial interactions (12, 13).
Morphologically, differentiation of the epididymis is characterized by growth and heavy coiling of the epithelial duct. In molecular and mechanistic terms, however, it is not completely understood which factors are involved in growth and differentiation of the Wolffian duct. We have recently described the effect of androgen on protein expression in a mouse fetal vas deferens (MFVD) cell line using a proteomics approach. Stimulation of MFVD cells with androgen resulted in posttranslational modification of two actin binding proteins, mElfin and CArG-binding factor A (CBF-A) (14). In the present investigation, we have applied proteomics to study and identify proteins that are involved in the androgen-dependent development of fetal rat epididymis and vas deferens, using freshly isolated or cultured tissues. Proteins regulated during fetal development were categorized as cytoskeletal proteins, nuclear proteins, transport proteins, chaperones, and glycolytic enzymes. Androgen regulation in vitro involved posttranslational modification of nuclear proteins and glycolytic enzymes, as well as RhoGDI1, a signaling molecule. Posttranslational modification of proteins by androgen stimulation may be a key regulatory event in genital tract development, suggesting a functional link between signaling pathways and cytoskeletal proteins.
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Materials and Methods |
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Tissue preparation
Rats were killed by CO2 gas and cervical dislocation. Fetuses were removed from the uteri and placed in PBS on ice. Epididymides and vasa deferentia were isolated as a whole from male fetal rats on E17, E18, E19, E19.5, E20, and E21. Dissection was performed in PBS in a sterile hood. All tissues were either immediately frozen on dry ice or fixed in Bouin’s solution. In addition, tissues from fetuses of E19 were used for organ culture studies. Epididymides, together with vasa deferentia, were placed in a drop of medium on a Millicell-CM 0.4-µm culture plate insert (Millipore Corp., Bedford, MA) that was floating on top of 0.5 ml medium (DMEM/F12 + 2% vol/vol charcoal-stripped fetal calf serum, 10 µg/ml insulin, 10 µg/ml transferrin, and a mixture of penicillin (100 IU/ml), streptomycin (100 µg/ml), and fungizone (0.6 µg/ml), all from Sigma (St. Louis, MO) in a four-well plate (Nunc, Roskilde, Denmark) (15). The two epididymides and vasa deferentia from one male were always cultured in either the absence of hormone (0.1% vol/vol ethanol vehicle) or in the presence of 10 nM of the synthetic androgen R1881, methyltrienolone (NEN Life Science Products, Boston, MA), for 9, 24, or 48 h. For each culture condition, 25 tissues were collected. For the frozen material, 40 tissues of E17 and E18 each and 25 tissues of E19-E21 were collected.
Two-dimensional gel electrophoresis (2DE)
All collected tissues from one time point were pooled and lysed in 500 µl 2D lysis buffer [7 M urea, 2 M thiourea, 4% wt/vol 3-[3-(cholamidopropyl)-dimethyl-ammonio]-1-propane sulfonate, 40 mM Tris-HCl (pH 8.8), 40 mM dithiothreitol (DTT) and 0.5% vol/vol immobilized pH gradient (IPG) buffer (pH 3–10)] using the Sample Grinding kit (Amersham Biosciences, Piscataway, NJ). High purity PlusOne chemicals were from Amersham Biosciences. Insoluble proteins and cell debris were pelleted at 100,000 rpm for 10 min at 4 C, using the Optima TLX Tabletop Ultracentrifuge (Beckman, Palo Alto, CA). Protein concentration was determined using the RC DC Protein Assay kit (Bio-Rad, Hercules, CA) and either 75 or 100 µg protein from total tissue lysate was used for isoelectric focusing (IEF). IEF was performed in an IPGphor apparatus according to the instructions of the manufacturer (Amersham Biosciences) using 24-cm nonlinear IPG strips (pH 3–10) (Amersham Biosciences). The strips were actively rehydrated overnight in rehydration buffer (8 M urea, 2% wt/vol 3-[3-(cholamidopropyl)-dimethyl-ammonio]-1-propane sulfonate, 40 mM DTT, and 0.5% vol/vol IPG buffer) containing the sample applying a voltage of 30 V/h (Vh). After rehydration, the strips were run until 100 kVh was reached. Following IEF, the strips were equilibrated in sodium dodecyl sulfate (SDS) buffer [6 M urea, 2% wt/vol SDS, 30% vol/vol glycerol, 50 mM Tris-HCl (pH 8.8)] containing 65 mM DTT for 15 min and thereafter in SDS buffer containing 135 mM iodoacetamide, for 2 min. 2DE was performed on 10% vol/vol Duracryl gels (Genomic Solutions, Perkin-Elmer, Boston, MA) using the Ettan Dalt gel caster and electrophoresis device (Amersham Biosciences) at a constant power of 3 W for 30 min and 180 W for 3–4 h, depending on the amount of gels run simultaneously, at 25 C. Gels were fixed overnight in 40% vol/vol methanol, 5% vol/vol ortho-phosphoric acid, and stained with colloidal Coomassie using a Colloidal Blue staining kit (Invitrogen, Paisley, Scotland, UK). Images were scanned with the Bio-Rad GS800 densitometer and analyzed using the PDQuest software package (Bio-Rad).
Mass spectrometry
Peptide samples were prepared as described previously (14). Briefly, proteins of interest were manually excised from the gel, destained twice with 30% vol/vol acetonitrile (ACN) in 50 mM ammoniumhydrogen carbonate for 15 min, dried in a Speed Vac Plus (Savant, NY) for 30 min and enzymatically digested overnight using sequencing grade trypsin (Promega, Madison, WI). Peptides were eluted from the gel with 30% vol/vol ACN/0.1% vol/vol trifluoric acid, and 0.5 µl peptide sample was mixed with 4 volumes matrix solution (2 mg -cyano hydroxycinnamic acid in 100% ACN), which was spotted on an AnchorChip target plate (Bruker Daltonik, Bremen, Germany). Mass spectra were generated on a Biflex III MALDI-TOF-MS (matrix-assisted laser desorption/ionization time-of-flight mass spectrometer) (Bruker Daltonik) and peptide fingerprints were analyzed using BioTools (Bruker Daltonik) and Mascot software (Matrix Science, London, UK) and the National Center for Biotechnology Information database.
Immunoprecipitation, SDS-PAGE, and Western immunoblotting
Immunoprecipitation was performed on 25 µg protein from total tissue lysate from rat E19 tissues cultured in the absence or presence of 10 nM R1881. Goat antirabbit-agarose beads were coupled to RhoGDI antibodies for 1 h at room temperature, cell lysate in PBS was added and the mixture rotated for 2 h at 4 C. The agarose-antibody precipitate was washed with PBS and the pellet was resolved in 25 µl Laemmli sample buffer for SDS-PAGE. Total tissue lysate (12.5 µg protein) or immunoprecipitate were loaded on 10% vol/vol and 15% vol/vol acrylamide gels. Gels were run on the mini PROTEAN gel system (Bio-Rad) and proteins were blotted to a nitrocellulose membrane (Schleicher & Schuell, Keene, NH). Western immunoblotting was performed using polyclonal antibodies against -fetoprotein (clone E19, 1:1000) and RhoGDI (clone A20, 1:1000) both from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Proteins were visualized by chemiluminescence detection (Western Lightning, Perkin-Elmer, Boston, MA).
Histology and immunohistochemistry
Tissues were fixed in Bouin’s solution for 24 h and were then washed in 70% vol/vol ethanol for 24 h. The fixed tissues were first embedded in 2% wt/vol agar before they were embedded in paraffin. Paraffin sections were cut at 7 µm and stained with hematoxylin/eosin. For immunohistochemistry, paraffin sections from E17 and E21 were incubated with an -fetoprotein antibody at a dilution of 1:500. Paraffin sections of E19 tissues cultured either in the absence or presence of R1881 for 48 h were used for immunohistochemistry with the RhoGDI antibody (1:500). Antibody incubations were performed overnight at 4 C. Secondary antibodies were coupled to peroxidase using the Strept ABComplex/HRP (Dako A/S Denmark), visualized with DAP stain (Pierce, Rockford, IL) and counterstained with hematoxylin.
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Results |
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. On E17, the epididymis is relatively small and underdeveloped. The epithelial duct consists of single layer of cuboidal cells with a small lumen, and no coiling of the duct is apparent (Fig. 1, A and C). On E21, the epididymis has grown much larger and the epithelial duct shows multiple coiling, which is apparent in the figure as multiple small cross-sections through the duct (Fig. 1D). It is obvious that marked tissue remodeling is needed to achieve such large changes in tissue structure, and this may require, and result in, differential expression and modification of a variety of proteins. To get more information about the spectrum and identity of proteins involved in growth-related tissue remodeling, we used a 2DE-driven proteomic approach, to identify proteins that are differentially regulated in time during this process of growth. Tissue lysates from the combined epididymides and vasa deferentia isolated on E17-E21 were used to separate proteins on 2D gels. All samples were run either in duplicate or triplicate. Protein expression profiles from E17-E21 were compared using the PDQuest software package.
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) representing at least 30 protein isoforms (Table 1). Analysis of the Coomassie-stained 2D gels, showed up-regulation of 36 protein isoforms (Fig. 2, red circles) and down-regulation of 17 protein isoforms (Fig. 2, green circles), which were consistently regulated throughout all duplicate gels and statistically significant. Inconsistently regulated proteins spots (e.g. doublet of spots 10 o’clock of spots 9) were excluded from the analysis. Furthermore, for three proteins we observed a shift in isoelectric point (pI) or other changes in one or more of the protein isoforms during development (Fig. 2, blue circles). These changes most probably represent posttranslational protein modifications.
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-fetoprotein precursor, AFP (spot 1, Fig. 2
). AFP occurred in the 2D gel as a 68-kDa multi-isoform protein and was down-regulated in time (Fig. 3A). However, two separate and fragmented isoforms of AFP were up-regulated in time. The up-regulation of the smaller AFP fragments was gradual but peaked at E19.5, after which the expression was going down again. The expression pattern of one of the AFP fragments is shown in Fig. 3D. Tropomyosin 5 was also up-regulated with a peak intensity at E19.5 (Fig. 3C). MALDI-TOF-MS revealed that fragmentation not only occurred for AFP but also for serum albumin precursor (SA), ß-actin, and -tubulin. Fragmentation of these proteins has been reported in relation to particular physiological roles (16, 17, 18). All the SA fragments were up-regulated gradually in time, of which one is illustrated in Fig. 3B.
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), and the regulation of AFP (spots 1 and 2, Fig. 2) and SA fragments (spots 4, Fig. 2), raised the question whether the tissue lysates contained serum proteins from infiltrating capillaries. To address this question, immunohistochemistry for AFP was performed on E17 and E21 tissues (Fig. 4, A and B). The staining shows that there is no detectable AFP in the epithelial duct, nor in the mesenchymal cells directly surrounding the duct, but there is positive staining in the cytoplasm of interstitial mesenchymal cells on E17. On E21, AFP staining in the interstitial mesenchyme was more abundant and some staining was also observed in the mesenchyme directly surrounding the epithelial duct. The staining was uniform and not restricted to capillaries, indicating that the tissue lysates contain serum proteins originating from the (interstitial) mesenchymal cells. Furthermore, no clear-cut difference in subcellular localization of the AFP staining between E17 and E21 tissue sections was visible. Western blot analysis of the tissue lysates showed a slight decrease of the two main bands of AFP (68 kDa) and an increase of some of the fragments, in agreement with the data of the 2D gels (Fig. 4C).
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) and microscopical (Fig. 5B) changes. After 9 h of culture, the epithelial duct is clearly visible as a lighter part of the tissues, and shows some minor coiling starting in the epididymis (Fig. 5A-1, 2). However, 48 h of culture in the absence of androgen resulted in regression of the epithelial duct, as demonstrated by the absence of the lighter duct in the tissue (Fig. 5A-3). In histological sections, some remnants of the epithelial duct are still present (Fig. 5B-3). Culturing the tissues in the presence of androgen for 48 h resulted in stabilization of the epithelial duct, and in most cases in increased coiling of the duct as a sign of differentiation (Fig. 5A-4). In histological sections, this coiling and stabilization is visible as cross-sections at three different positions (Fig. 5B-4). Comparing the sections of Figs. 1B and 5B-4 illustrates that development of the epididymis in vitro during a culture period of 48 h, although much slower, occurs in a similar pattern as in vivo. Furthermore, we also noticed some swelling of the tissues in culture, irrespective of the presence or absence of androgen.
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spots a–c and e, and Table 1], and one protein was visibly up-regulated (hnRNPA3; Fig. 6A, spot d). Comparison of these results with Fig. 2 and Table 1 showed that TPI, G3PD, hnRNP A2/B1, and hnRNP A3 were also regulated in the in vivo situation.
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). One protein isoform of RhoGDI1 was identified in E19 tissues that were either directly frozen or cultured. However, culturing the tissues in the presence of androgen for 48 h resulted in the appearance of a second, more acidic, protein isoform that was also identified by MALDI-TOF-MS as RhoGDI1. The glycolytic enzyme TPI was regulated by R1881 in vitro as well as in vivo at the level of posttranslational modification (Fig. 6C). In the presence of androgen (E19, t = 0 and +10 nM R1881 for 9 h and 48 h), expression of a more basic isoform was relatively high, although the intensity decreased with increasing culture time. Androgen withdrawal resulted in a further decrease in the expression of the basic isoform, whereas a more acidic isoform appeared (Fig. 6, C and D). Apparently, androgens play a role in maintaining TPI in a more basic isoform.
RhoGDI1 is an interesting candidate protein to analyze in more depth because it was reported that this protein can increase transcriptional activity of the AR (19). We questioned whether addition of androgen resulted in a change in subcellular localization of RhoGDI1 and whether the second isoform represented a differential phosphorylated isoform because phosphorylation of RhoGDI1 has been reported (20). Immunohistochemical analysis of epididymis cultured in the absence of androgen for 48 h, confirmed the cytoplasmic localization that was already reported (21). As expected, no nuclear localization of RhoGDI1 was observed after androgen treatment (Fig. 7, A and C and B and D). Although the epithelial duct regresses in the absence of R1881, RhoGDI1 is still expressed in the cytoplasm of the remnants of the duct, and also in the surrounding mesenchyme. Western immunoblot analysis of tissue lysates showed an equal expression of RhoGDI under all culture conditions (Fig. 7E). Immunoprecipitation of RhoGDI1 from the lysates with a RhoGDI1 antibody and subsequent immunoblot analysis with a phospho-serine antibody confirmed that RhoGDI1 is a phosphoprotein (Fig. 7F).
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Discussion |
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Identification of growth-related proteins
Using 2DE driven proteomics, 20 proteins were identified that are regulated during fetal growth and development of the rat epididymis and vas deferens. The proteins identified can be categorized into enzymes (-enolase, TPI, G3PD, probable thioredoxin peroxidase 2), cytoskeletal proteins (-tubulin, ß-actin, tropomyosin 5, F-actin capping protein -2), transport proteins (AFP, SA, transferrin, fibrinogen ß/
), RNA binding proteins (hnRNP A2/B1/A3), and chaperones (Hsp-47 precursor). Furthermore, fragmented isoforms of SA, AFP, -tubulin, and ß-actin were identified, which were up-regulated in time during genital tract development.
AFP is a specific fetal glycoprotein that is secreted by embryonic tissues and is involved in up- and down-regulation of cell growth (22), in immunosuppression, and in apoptosis (23). AFP belongs to the proteins encoded by the serum albumin multigene family, which present three structurally homologous domains I, II, and III (16). Fragmentation of AFP has been reported, with cleavage products of 32 and 38 kDa resulting from domains I and III. These products can be further fragmented into proteolytically stable isoforms of 23 and 26 kDa (17). A similar kind of fragmentation was described for SA, but with many more proteolytic cleavage products possible (24). These reported fragments of AFP correspond in size to the fragments that were identified in the present study. Furthermore, others have described that distinct AFP isoforms accumulate at different stages of fetal development (25, 26) and that AFP can interact with nuclear receptors (27). Thus, the up-regulation of AFP fragmented isoforms may be physiologically relevant for growth and differentiation of epididymis and vas deferens.
Glycolytic enzymes interact directly (-enolase, G3PD), or indirectly (TPI), with structural proteins and complexes like actin and microtubules (28, 29). It was recently described for TPI that it binds to the actin-binding protein cofilin, which in turn binds to Na,K-ATPase upon phosphorylation by the 
-signaling pathway (30). TPI takes part in production of energy, which is transduced to the cytoskeleton that modulates cell function, proliferation, and differentiation (30). The cytoskeletal proteins actin and tubulin, and in particular specific cleavage of these proteins, have been implicated in several regulatory processes, including apoptosis (18) and other processes such as meiosis resumption in starfish oocytes (30) Our finding that fragments of ß-actin and -tubulin accumulate taken together with the observation of posttranslational regulation of glycolytic enzymes could reflect the processes of vigorous tissue remodeling during the process of genital tract development.
Identification of androgen-regulated proteins during growth
Because it is known that androgen action is essential for growth and development of the male genital tract, it was our aim to identify androgen-regulated proteins that are involved in this process. Epididymides and vasa deferentia cultured in vitro in the absence and presence of androgen were subjected to 2DE, and the differentially expressed proteins identified were compared with the proteins identified in vivo. Glycolytic enzymes G3PD and TPI, and nuclear proteins hnRNP A2/B1/A3 were identified to be regulated by androgen, as was also observed in vivo. In addition, RhoGDI1 was identified as a protein that was regulated by androgen at the level of posttranslational modification. Although the regulation of these proteins occurs under the action of androgen, we cannot conclude that this is the result of a direct and/or indirect androgen action. Furthermore, we did not observe any obvious regulation of the AR protein in the 2D gels. However, the AR protein is of lower abundance and therefore not easily detectable in such a broad range 2D gel.
Rho guanine nucleotide dissociation inhibitor 1(RhoGDI1 or RhoGDI), is a cytoplasmic protein originally identified as a negative regulator of RhoGTPases (21). RhoGTPases are molecular switches that cycle between an active membrane-associated GTP-bound state and an inactive cytoplasmic GDP-bound state. This switch is carefully controlled by exchange factors, activating proteins, and dissociation inhibitors (31). RhoA, Rac, and Cdc42 belong to the family of RhoGTPases, which regulate many signal transduction pathways, including those linked to the actin cytoskeleton, microtubule dynamics, vesicular transport dynamics, regulation of cell polarity, gene transcription, G1 cell cycle progression, and a variety of enzymatic activities (31, 32). Members of the RhoGDI family block GDP dissociation from RhoGTPases and control cytoplasmic localization of RhoGTPases. In addition, it was described that RhoGDI1 specifically increases the transcriptional activity of estrogen receptors (ER, ERß), glucocorticoid receptor and AR (19). This activation is mediated via repression of RhoGTPases, which demonstrates that the -mediated signaling pathway is an important regulator of ERs, glucocorticoid receptor, and AR transcriptional activity (19). Although RhoGDI1 is ubiquitously expressed in all tissues, RhoGDI1 knockout mice show specific and progressive impairment of kidneys and reproductive organs probably as a result of the destruction of the actin cytoskeleton (33), meaning that the function of RhoGDI1 in the kidney and reproductive organs is nonredundant.
The testis of RhoGDI1 -/- mice reveals structural abnormalities, the number of germ cells is dramatically decreased and mature sperm cells are only rarely detected in seminiferous tubules and epididymides (33). RhoGDI1 -/- female mice have an intrinsic defect in their reproductive system, which is most evident in the postimplantation development of RhoGDI1 -/- embryos. For the epididymis, no functional developmental defect in the knockout mice was reported (33).
Bourmeyster et al. (20) have detected RhoGDI1 on a 2D gel as a single protein spot with a pI of 5.1, which represents the free, unbound form of RhoGDI. In the case were RhoGDI was associated with RhoA GTPase, they detected two RhoGDI protein spots with pI 4.6–5.1, which could be reduced to one protein spot after phosphatase treatment (20). Furthermore, they have shown that RhoA-RhoGDI1 association depends on RhoGDI1 phosphorylation. Thus, phosphorylation and dephosphorylation of RhoGDI1 determines the activation state of RhoA GTPase (20). In the present tissue culture experiments, 48 h of androgen treatment resulted in the appearance of a second, more acidic, RhoGDI1 isoform that could represent a phosphorylated isoform. If RhoGDI1 is indeed phosphorylated by androgens, this could lead to complex formation with Rho GPTase and repression of the Rho GTPase activity. Taken the fact that RhoGDI1 enhancement of AR transcriptional activity occurs via RhoGTPase repression (19), one could postulate that RhoGDI1 phosphorylation leads to enhanced AR transcription activation, and as a result regulation of RhoGDI1 activity by phosphorylation and dephosphorylation might function as a feedback loop in androgen signaling.
hnRNPs form a large family of proteins that are categorized on the basis of structural/functional motifs, and of which the A and/or B type are the most abundant hnRNP proteins (34). Besides a large number of posttranscriptional isoforms, the hnRNP A and/or B proteins also show extensive posttranslational modifications. For a specific subset of hnRNPs, hnRNP A2, hnRNP A3, and hnRNP-related DNA binding protein 40/CBF-A, an interaction with nuclear actin was described, and an interaction was suggested for cytoplasmic actin (34). In a previous study, we have identified CBF-A, which is a minor variant of hnRNP A2, to be regulated by androgen at the level of posttranslational modification in the mouse fetal vas deferens cell line MFVD (14). This coincides with the present finding that androgens regulate hnRNPs posttranslational modification in genital tract tissues.
Glycolytic enzymes, like TPI and G3PD, are dependent on androgen action for their activity, as was demonstrated by castration experiments. Enzymatic activity was found to be decreased in rat epididymis and monkey seminal vesicle after castration, which was restored after androgen replacement (35, 36, 37). Our present results suggest that androgenic regulation of glycolytic enzyme activity is controlled at the level of posttranslational modification.
In summary, our study shows that changes in protein expression profiles can be detected on 2D gels, when developing genital tract tissues are followed in time in vivo. Analyses of androgen-regulated proteins in epididymis and vas deferens tissues exposed to androgen, also revealed several changes in protein expression profiles when grown in vitro. In the present system, androgen action occurs mainly at the level of posttranslational modification. We hypothesize that during the process of androgen stimulation, which will lead to growth and differentiation of the male genital tract, posttranslational modification of glycolytic enzymes regulates their activity and their association with cytoskeletal proteins. Furthermore, androgen-induced posttranslational modification of RhoGDI1 and glycolytic enzymes may be an important functional link between signaling pathways and cytoskeletal rearrangements in control of growth and development of the male internal genital tract.
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Acknowledgments |
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Footnotes |
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-fetoprotein; AIS, androgen insensitivity; AR, androgen receptor; 2D, two-dimensional; 2DE, 2D gel electrophoresis; DHT, dihydrotestosterone; DTT, dithiothreitol; E, embryonic day; ER, estrogen receptor; G3PD, glyceraldehyde 3-phosphate dehydrogenase; hnRNP, heterogeneous nuclear ribonucleoprotein; IEF, isoelectric focusing; IPG, immobilized pH gradient; MALDI-TOF-MS, matrix-assisted laser desorption/ionization time-of-flight mass spectrometer; MFVD, mouse fetal vas deferens; pI, isoelectric point; SA, serum albumin precursor; SDS, sodium dodecyl sulfate; T, testosterone; TPI, triosephosphate isomerase. Received March 31, 2003.
Accepted for publication June 11, 2003.
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References |
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