Skip to main content
NCBI home page
Endocrinology logoLink to Endocrinology
. 2008 Feb 21;149(6):2959–2969. doi: 10.1210/en.2007-1078

Androgen Receptor Is Causally Involved in the Homeostasis of the Human Prostate Endothelial Cell

Alejandro Godoy 1, Anica Watts 1, Paula Sotomayor 1, Viviana P Montecinos 1, Wendy J Huss 1, Sergio A Onate 1, Gary J Smith 1
PMCID: PMC2408806  PMID: 18292195

Abstract

Androgen deprivation causes a reduction of blood flow in the prostate gland that precedes temporally apoptosis of the epithelium. The acute response of prostate endothelial cells to androgen deprivation suggested they represent a primary target for androgen. However, rat prostate endothelial cells were reported not to express androgen receptor (AR), and the role of the androgen axis in human prostate endothelial cell (HPEC) homeostasis was poorly characterized. In this study AR expression was detected in HPEC in vivo in clinical specimens of benign prostate and prostate cancer, and AR function as a transcription factor was demonstrated in HPEC in primary xenografts of human benign prostate tissue transplanted into severe combined immunodeficient mice by iv administration of adenoviral mouse mammary tumor virus-driven luciferase expression vector. AR expression and functionality were maintained in vitro in primary cultures of HPEC that coexpressed CD31, CD34, von Willebrand factor, intercellular adhesion molecule, vascular endothelial growth factor receptor 1, and vascular endothelial growth factor receptor 2 but did not express prostate-specific antigen. AR expression in primary cultures of HPEC isolated from surgical specimens of benign prostate was validated using RT-PCR, cDNA sequencing, immunocytochemistry, and Western blot analyses. Scatchard analyses demonstrated a single ligand-binding site for R1881 in primary cultures of HPEC, with dissociation constant of 0.25 nm, and AR-mediated transcriptional activity was demonstrated using adenoviral mouse mammary tumor virus-driven luciferase reporters. Dihydrotestosterone increased proliferation in primary cultures of HPEC in a dose-dependent manner without modulating endothelial tube formation in Matrigel (BD Biosciences, Bedford, MA). Therefore, HPECs express functional AR, and androgen plays a direct role in modulating HPEC biology.

HUGGINS AND HODGES (1) reported in 1941 that growth of prostate cancer (CaP) depended on androgen, and this conceptual breakthrough led to the development of androgen deprivation therapy (ADT), the standard treatment for advanced CaP for over 60 yr. ADT reduced the level of circulating testicular androgens and inhibited the stimulatory effect of androgen on CaP (2). Consistent with the observed reduction in CaP mass and growth rate, the primary target for ADT in prostate tissue was presumed to be the epithelial cell compartment. Androgens regulate prostate epithelial cells directly, and indirectly through stimulation of prostate stromal cells to produce autocrine and paracrine-acting growth and differentiation factors, during organogenesis and in the adult, as well as in CaP (3,4,5). However, ADT is rarely curative, and the initial response to ADT is followed, in virtually all cases, by relapse of the disease as hormone-refractory CaP, the lethal phenotype of the disease (6).

Ten years ago, two groups reported that the initial observable physiological effect of androgen deprivation on the rat prostate gland was a significant reduction in blood flow (7,8). The effect of castration on blood flow was observed in ventral prostate, but not in dorsal prostate or in the Dunning R3327 prostate tumor xenograft model (8). Perturbation of the prostatic vasculature was evident as early as 18 h after castration, and the decreased blood flow in the rat ventral prostate was correlated with the appearance of apoptotic endothelial cells (7,9). Because the appearance of apoptotic endothelial cells preceded the appearance of apoptotic epithelial cells by several days, both groups hypothesized that a large proportion of prostate epithelial cell loss was an indirect effect caused by hypoxic/ischemic conditions within the prostate gland that resulted from castration-induced endothelial cell death and reduction in blood flow.

Rat prostate endothelial cells were reported to lack expression of androgen receptor (AR) (10). Therefore, it was anticipated that an androgen-regulated intermediary paracrine molecule, perhaps a growth factor synthesized by AR-expressing prostate epithelial or stromal cells, regulated survival of prostate endothelial cells (11,12). In support of this hypothesis, castration of severe combined immunodeficient (SCID) mice transplanted with the androgen-dependent Shionogi carcinoma demonstrated that involution of tumor vessels was concomitant with decreased vascular endothelial growth factor (VEGF) expression in tumor epithelial cells (12). However, AR expression was observed in human endothelial cells from several tissues, including skin (13,14), salivary gland (15), bone (16), bone marrow (17), corpus cavernosum from the penis (18), and most recently, skeletal muscle (19). In prostate tissue, El-Alfy et al. (20) observed immunohistochemical staining for AR of several nuclei of human prostate endothelial cells (HPECs), which suggested the potential of a direct role for androgens in modulating HPEC biology. However, expression of AR in HPECs was controversial, and functional activity of AR has not been demonstrated directly in HPECs.

This study analyzed the expression and functionality of AR in HPECs in vivo and in vitro. AR expression was observed in vivo in histological specimens of human benign prostate and CaP at comparable levels of intensity. Primary cultures of HPECs and primary xenografts of human benign prostate tissue maintained expression of functional, high-affinity AR that transactivated mouse mammary tumor virus (MMTV) promoter-driven luciferase gene reporter constructs in the presence of dihydrotestosterone (DHT). Moreover, DHT, through AR, directly increased proliferation of primary cultures of HPECs in a dose-dependent manner without affecting formation of endothelial tube structures in Matrigel (BD Biosciences, Bedford, MA), suggesting that the differentiation and migration processes involved in endothelial tube formation are independent of proliferation in prostate endothelial cells. These studies provide the first evidence that androgens, through AR, play a direct role in modulation of HPEC biology and suggest that the human prostate vasculature might represent a primary therapeutic target for ADT.

Materials and Methods

Human prostate primary xenografts and in vivo adenoviral infection

Surgically resected prostate tissue was collected in accordance with National Institutes of Health guidelines on the use of human subjects and with the approval of the Internal Research Board at Roswell Park Cancer Institute (RPCI). Benign human prostate tissue was obtained from patients during radical prostatectomy or cystoprostatectomy. Tissue specimens were collected and submerged immediately in ice-cold ViaSpan solution (Barr Laboratories Inc., Pomona, NY) and transported on ice for transplantation.

Primary xenografts of human prostate tissues were established in SCID mice as described previously (21,22). All animal experimentation was conducted in accordance with the guidelines of the National Institutes of Health and the Institutional Animal Care and Use Committee at RPCI. In brief, tissue specimens were minced into 8-mm3 pieces, and the pieces were transplanted immediately into 2- to 3-month-old SCID mice that were previously implanted sc with 12.5 mg sustained-release testosterone pellets (Innovative Research of America, Sarasota, FL) to maintain a serum testosterone level of approximately 4.0 ng/ml throughout the study. For implantation of prostate tissue, small (∼3 mm) incisions were made in the skin on the right and left flanks of mice anesthetized with Domitor (1 mg/kg ip; Pfizer, Inc., New York, NY). Tissue pieces to be implanted were dipped in Matrigel, and the coated tissue pieces were inserted into the sc space through a 10-gauge trocar device (Popper & Sons. Inc., Lincoln, RI). Between three and five tissue pieces from a single patient were implanted individually along each flank. Incision sites were closed with Nexband tissue glue (Veterinary Products Laboratories, Phoenix, AZ).

Infection of endothelial cells in the xenografts with adenovirus containing AR-driven reporter constructs was initiated at d 14 after implantation. Briefly, SCID mice transplanted with human benign prostate tissue were injected iv with 300 μl of a saline solution containing an adenoviral expression vector encoding an MMTV or prostate-specific antigen (PSA) promoter-driven luciferase reporter [∼108 infectious units (ifu)/ml, BD Adeno-X Virus Purification and Rapid Titer Kits; BD Biosciences, Bedford, MA]. As a control, SCID mice transplanted with human benign prostate tissue were injected iv with 300 μl of a saline solution containing an adenoviral expression vector encoding green fluorescence protein (GFP) under control of cytomegalovirus (CMV) promoter (∼108 ifu/ml), an expression construct independent of AR transactivation. After 48-h infection, host animals were euthanized, and xenograft tissues were harvested, fixed in 10% formalin for a minimum of 24 h, and embedded in paraffin for histological analysis. Paraffin blocks were sectioned (5 μm) onto ProbeOn Plus slides (Fisher Scientific Intl., Suwannee, GA) and tissue sections processed for standard immunohistochemical procedures, as described below. Luciferase and GFP expression in endothelial cells was demonstrated using the polyclonal antibodies goat anti-Luciferase (1:2000, CHEMICON International Inc., Temecula, CA) and rabbit anti-GFP ab-6556 (1:1000, Abcam, Inc., Cambridge, MA), respectively. Glucocorticoid receptor (GR) and progesterone receptor (PR) expression in HPECs was analyzed using the polyclonal antibodies rabbit anti-GR M-20 (1:200; Santa Cruz Biotechnology, Inc., Santa Cruz, CA) and rabbit anti-PR (1:250; Dako Corp., Carpinteria, CA), respectively.

Prostate tissue digestion and primary cell culture

Primary cultures of HPECs were prepared using a Dynabeads-based methodology (DYNAL Biotech ASA, Oslo, Norway) (23). Surgical specimens of prostate tissue were minced in 8-mm3 pieces and digested with dispase (2.4 U/ml; Invitrogen Corp., Carlsbad, CA) for 16 h at 4 C. The heterogeneous tissue hydrolysate was cultured for approximately 1 wk in endothelial growth media (Endothelial cell growth medium MV, Promocell, Heidelberg, Germany) supplemented with 5% fetal calf serum at 37 C in 5% CO2. After 1 wk in culture, HPECs were isolated using anti-CD31-conjugated magnetic beads, and cultured in endothelial growth media for no more than six passages before experimental studies were performed. Purity of endothelial cell cultures was estimated by staining the primary cultures with an anti-von Willebrand factor antibody, with more than 95% of the cells expressing von Willebrand factor in culture.

Immunostaining

Tissue sections of formalin-fixed and paraffin-embedded human benign prostate and CaP tissue were obtained from 10 randomly selected clinical cases provided by the Department of Pathology at RPCI. Primary cultures of HPECs, human umbilical vein endothelial cells (HUVECs), or the LNCaP human CaP cell line were fixed in situ using 4% (wt/vol) paraformaldehyde for 30 min at room temperature. Before immunostaining, endogenous peroxidase activity was inhibited with 0.3% (vol/vol) H2O2 in methanol, and nonspecific binding of antibodies was blocked with 3% (wt/vol) BSA (EMD Chemicals, Gibbstown, NJ) for 30 min at room temperature. Histological sections and fixed cells were immunostained using standard methodology (24,25,26). AR and CD31 colocalization analyses were performed using the mouse/rabbit EnVision G/2 Double Stain System, according to manufacturer’s instructions (Dako). Tissue sections or cells were incubated overnight with mouse anti-CD31 (1:20; Dako), mouse anti-CD34 (1:50; Neomarkers, Fremont, CA), mouse anti-von Willebrand (1:10; Biogenex, San Ramon, CA), and rabbit anti-AR (N-20) antibody (1:100; Santa Cruz Biotechnology). For proliferation assays, cells were incubated overnight with mouse anti-Ki-67 (1:1000; Vision BioSystems Inc., Norwell, MA) antibody. All antibodies were diluted in 100 mm Tris-HCl buffer (pH 7.8) that contained 8.4 mm sodium phosphate, 3.5 mm potassium phosphate, 120 mm NaCl, and 1% (wt/vol) BSA. After washing three times in Tris-HCl buffer (pH 7.8) for 10 min each, specimens were incubated with horseradish peroxidase (HRP)-conjugated antirabbit IgG or antimouse IgG (1:100; Dako) for 2 h at room temperature. Peroxidase activity was developed using 100 mm Tris-HCl buffer that contained 3,3-diaminobenzidine tetrahydrochloride (1 μg/ml; Sigma-Aldrich, St. Louis, MO) and H2O2 (1 μl/ml; VWR Intl., West Chester, PA). Colocalization analyses of AR/GR/PR with CD31 used the mouse/rabbit EnVision G/2 Double Stain System, with AR/GR/PR expression detected using a HRP-conjugated secondary antibody and 3,3-diaminobenzidine tetrahydrochloride substrate (brown precipitate) and CD31 expression visualized using an alkaline phosphatase-conjugated secondary antibody and Permanent Red substrate (red precipitate). Hematoxylin was used as a nuclear counterstain in tissue sections. Stained slides were dehydrated by sequential steps through a graded series of alcohol washes and Citrisolv (Fisher Scientific International, Suwannee, GA), and mounted using coverslips. For immunofluorescence studies, cells were incubated for 2 h with Cy2- or Cy3-conjugated affinity purified donkey antirabbit or antimouse IgG (1:200; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA) at room temperature. 4′, 6′-diamidino-2-phenylindole dihydrochloride was used as the counterstain to visualize nuclei. Immunocytochemistry in the absence of primary antibody, or using preimmune serum, provided negative controls.

Endothelial cell tube formation assay

Matrigel (176 μl) was dispensed to each well of a 24-well plate and incubated at 37 C in 5% CO2 for 30 min for the Matrigel to solidify. HPECs were suspended at a density of 100 × 103 cells per ml in 500 μl serum-supplemented endothelial cell growth media (Promocell), or endothelial cell growth media supplemented with 5% (vol/vol) charcoal-stripped fetal bovine serum (FBS) (HighClone Laboratories, Logon, UT), and designated concentrations of DHT (0.01, 0.1, 1, and 10 nm). HPECs (50 × 103 cells) were seeded into wells containing Matrigel and incubated for 12 h at 37 C in 5% CO2. The effect of DHT on endothelial cell tube formation was analyzed by collecting four random digital images (10× magnification) from each well, and total tube length per image was quantified using Optimas 6.2 (Media Cybernetics, Inc., Bethesda, MD). The total length of tubular structures per image was calculated by the software, which assigned length in arbitrary units. The total tube length per digital image was a summation of the lengths of all individual tubular structures.

RT-PCR

Total RNA from primary cultures of HPECs, HUVECs, and LNCaP cells was prepared using the RNAeasy mini-kit (QIAGEN, Inc., Valencia, CA). RT from mRNA was performed using the SuperScript III First-Strand kit (Invitrogen) (24). Approximately 1 μl reverse transcribed cDNA product was used as template in the Platinum PCR Supermix (Invitrogen) reaction mix that contained 200 nm each primer. PCR products were separated using electrophoresis in 2% agarose gels and visualized with ethidium bromide. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a loading control in the analytical gels. The identity of the specific PCR product was confirmed by sequencing the product (RPCI core facility). Primer sequences for the PCRs were: CD31, forward 5′-ATGATGCCCAGTTTGAGGTC-3′ and reverse 5′-ACGTCTTCAGTGGGGTTGTC-3′; CD34, forward 5′-CCTCTCCA-GAAAGCTGAACG-3′ and reverse 5′-GTGGGGGATTCTTGCTTTTT-3′; von Willebrand, forward 5′-TGCTGTGACACATGTGAGGA-3′ and reverse 5′-AGAAGGGCACAAGAGCAGAA-3′; ICAM, forward 5′-GAACATAGGTCTCTGGCCTCAC-3′ and reverse 5′-ATTGTGAACA-CTGGCAGAAATG-3′; VEGFR1, forward 5′-TTTGGATGAGCAGTGTGAGC-3′ and reverse 5′-GTGCTGCATCCTTGTTGAGA-3′; VEGFR2, forward 5′-AAACCCATACCCTTGTGAAGAAT-3′ and reverse 5′-CA-AACGTAGATCTGTCTGCAGTG-3′; PSA, forward 5′-CTCTACGATATGAGCCTCCTGAA-3′ and reverse 5′-AGGTCCATGACCTTCACAGC-3′; AR, forward 5′-TACCAGCTCACCAAGCTCCT-3′ and reverse 5′-GC-TTCACTGGGTGTGGAAAT-3′; and GAPDH, forward 5′-GTCTTCACCACCATGGAGAAG-3′ and reverse 5′-CAAAGTTGTCATGGATG-ACCTTGG-3′.

Preparation of nuclear extracts

Nuclear extracts from primary cultures of HPECs and LNCaP cells were obtained using the NE-PER Nuclear and Cytoplasmic Extraction Kit (Pierce, Rockford, IL). Briefly, a 20 μl packed cell volume (∼40 mg protein) was lysed for 10 min with 200 μl CERI buffer (Pierce) and 1 min with 11 μl CERII buffer (Pierce). Lysates were centrifuged for 5 min at approximately 16,000 × g at 4 C, and the supernatant (cytoplasmic fraction) was transferred to a clean prechilled tube and stored at −80 C. The insoluble fraction (pellet) that contained the nuclei was resuspended in 100 μl ice-cold NER buffer (Pierce), incubated for 40 min on ice, and centrifuged at approximately 16,000 × g at 4 C.

Immunoblotting

Nuclear extracts (30 μg protein) were separated electrophoretically using SDS-PAGE (10% wt/vol; Bio-Rad Laboratories, Inc., Hercules, CA) under reducing conditions and the separated proteins transferred to nitrocellulose membranes using standard procedures (24). Membranes were blocked in 5% (wt/vol) nonfat dry milk in Tris-buffered saline (TBS) [10 mm Tris-HCl (pH 7.4) and 150 mm NaCl] for 30 min at room temperature and were incubated overnight with primary antibody diluted in 5% (wt/vol) nonfat dry milk in TBS. Primary antibodies were rabbit anti-AR (1:1000; Santa Cruz Biotechnology) and rabbit anti-topoisomerase I (1:500; Santa Cruz Biotechnology). Membranes were washed twice with Tris-HCl [Tris-buffered saline with Tween 20 (pH 7.4)] containing 2.5 m NaCl, 0.05% (vol/vol) Tween 20, and 0.2% (vol/vol) Triton X-100, and incubated with an appropriate amount of HRP-conjugated secondary antibody diluted in 5% (wt/vol) nonfat dry milk in TBS, for 2 h at room temperature. Antibody localization was visualized using enhanced chemiluminescence (Pierce). Equal loading of nuclear extract protein samples and success of membrane transfer were verified visually by staining membranes with ponceau solution, or anti-topoisomerase I antibody (Santa Cruz Biotechnology).

AR-ligand binding assays

LNCaP and primary cultures of HPECs were preincubated for 12 h in RPMI 1640 or endothelial cell growth media (Promocell) containing 5% (vol/vol) charcoal-stripped FBS before binding assays with radiolabeled steroid ligand. Total R1881 binding to AR was determined by incubating cells in increasing concentrations of [17∝-methyl-3H]R1881 (PerkinElmer, Inc., Wellesley, MA), ranging from 0–6.0 nm, for 4 h at 37 C. Nonspecific binding was determined by performing total binding analyses in the presence of a 500-fold excess of nonradioactive R1881 (PerkinElmer) under the same experimental conditions. At the end of the incubation, cells were washed three times with ice-cold PBS, bound R1881 extracted in ethanol, and the extracted radiolabeled steroid measured using scintillation spectrometry (LS 6500 Multi-Purpose Scintillation Counter; Beckman Coulter, Inc., Fullerton, CA). Specific binding of [3H]R1881 was calculated by subtraction of the nonspecifically bound radioactivity from the total bound radioactivity. The dissociation constant (Kd) for R1881 and the number of binding sites were determined using Scatchard analysis (27). The Kd value was represented by the negative reciprocal of the slope (−1/Kd), and the number of binding sites was calculated based on the maximum binding capacity (maximum binding capacity × intercept) and Avogadro’s number, assuming a 1:1 R1881/AR binding stoichiometry. The Kd value is represented two independent experiments performed in triplicate.

Luciferase assays

LNCaP, CV-1, and primary cultures of HPECs grown in 24-well plates were washed, and incubated for 12 h in RPMI 1640, DMEM, or endothelial cell growth media, respectively, containing 5% (vol/vol) charcoal-stripped FBS. Cells were infected for 3 h with an adenoviral expression vector encoding an MMTV promoter- or PSA promoter-driven luciferase reporter (multiplicity of infection between 10 and 20 ifu/cell). After infection, cells were incubated 36 h in the corresponding media containing 5% (vol/vol) charcoal-stripped FBS and supplemented with the designated concentration of DHT. For demonstration of inhibition of MMTV-driven luciferase reporter activity by the antiandrogen flutamide, cultures of HPECs were maintained in the presence of designated concentrations of flutamide (0.2, 2, 20, and 200 μm) during the entire experiment (3-h infection and 36-h incubation). At the conclusion of the incubation, cells were washed with PBS and lysed in 50 μl reporter lysis buffer (Promega, Madison, WI), followed by a freeze-and-thaw incubation to ensure lysis. Lysates (25 μl) were assayed for luciferase activity using the Luciferase Assay System (Promega) and Veritas microplate luminometer (Turner BioSystems, Sunnyvale, CA). Luciferase values were normalized to total protein content measured using the Bradford assay (Bio-Rad Laboratories). All experiments were performed in triplicate.

Statistical analysis

Statistical evaluation of the data was performed using SuperANOVA software (Abacus Concepts, Berkeley, CA). All data differences were considered statistically significant when the P value was less than 0.05. Error bars on graphs represent sd.

Results

HPECs express functional AR in vivo

The expression of AR in HPECs in tissue sections of human benign prostate and CaP was demonstrated by AR and CD31 colocalization analyses using the mouse/rabbit EnVision G/2 Double Stain System. In human benign prostate specimens, AR was localized to the nucleus of almost all luminal epithelial cells, and to nuclei of some basal epithelial cells (Fig. 1, A and C). In the stromal compartment, nuclei of some fibromuscular cells immunostained for AR. However, unstained nuclei of fibromuscular cells also were observed (Fig. 1, A–C). In blood vessels of benign human prostate tissue, the nuclei of endothelial cells lining the vascular lumens demonstrated heterogeneous immunostaining for AR, whereas the majority of endothelial cells showed positive AR immunostaining (Fig. 1, B and C, black arrows) some endothelial cells were not immunoreactive (Fig. 1, B and C, blue arrows). No differences in the level of intensity of AR immunostaining were observed between endothelial cells in the microvasculature and macrovasculature (data not shown). In CaP tissue specimens across a diversity of Gleason grades, AR immunostaining was observed in almost all nuclei of epithelial cells (Fig. 1D), and in some nuclei of fibromuscular cells (Fig. 1, D and E). Nuclei of most endothelial cells in CaP immunostained for AR (Fig. 1, D and E), as observed for endothelial cells in benign human prostate. No differences in the level of intensity of AR immunostaining were observed between endothelial cells in the microvasculature and macrovasculature of CaP (data not shown).

Figure 1.

Figure 1

Open in a new tab

AR and CD31 double immunostaining in clinical specimens of human benign prostate and CaP. AR and CD31 colocalization studies were performed using the mouse/rabbit EnVision G/2 Double Stain System. AR expression was visualized using 3,3-diaminobenzidine tetrahydrochloride substrate (brown staining), and CD31 expression was visualized using a Permanent Red substrate (red staining). A–C, In human benign prostate, AR expression was localized predominantly in the nuclei of luminal epithelial cells (A and C), stromal cells (A–C), and endothelial cells (A–C, black arrows). However, some endothelial cells did not immunostain for AR (B and C, blue arrows). D and E, In CaP, immunostaining for AR was observed in cancer epithelial cells (D), stromal cells (D and E), and endothelial cells (D and E, black arrows). Similar to benign prostate, a subset of endothelial cells was not immunoreactive for AR. Bars, 10 μm.

In vivo functional activity of AR in HPECs was characterized in primary xenografts of benign human prostate tissue that retain a human vasculature that maintains a marked angiogenesis capability (21,22). SCID mice with primary xenografts of human benign prostate tissue were injected iv with a saline solution containing an adenoviral MMTV- or PSA-driven luciferase gene reporter. After 48 h of infection, xenografts were harvested, fixed, and embedded in paraffin. Luciferase expression in the human endothelial cells was detected using immunocytochemistry (Fig. 2). Endothelial cells in xenografts of benign prostate tissue harvested from mice infected with the MMTV-driven luciferase reporter demonstrated AR-mediated cytoplasmic expression of luciferase (Fig. 2A). Some stromal cells close to vessels also immunostained for luciferase (Fig. 2A, blue arrows). In contrast, luciferase expression was not detected in any endothelial cells in mice infected with a PSA-driven luciferase reporter (Fig. 2B), a reporter construct specific for AR-mediated transactivation in prostate epithelial cells. As a control for infection efficiency, mice were infected with an adenovirus containing a CMV-driven GFP construct that is expressed in all cells independent of AR. High efficiency of infection with GFP adenovirus was observed in human endothelial cells in the xenografts of human benign prostate tissue, with most endothelial cells expressing GFP after 48-h infection (Fig. 2C).

Figure 2.

Figure 2

Open in a new tab

In vivo functional activity of AR in HPECs. SCID mice previously transplanted with 8-mm3 pieces of freshly harvested human benign prostate tissue, and the engrafted tissue allowed 14–30 d for establishment, were injected iv with an adenoviral (Ad) MMTV promoter (A) or PSA promoter (B) driven luciferase gene reporter, or an adenoviral GFP reporter under the control of a CMV promoter (C). In mice injected with the MMTV adenoviral construct, immunoreaction for luciferase was observed in endothelial cells of the primary xenografts (A, black arrows), but also in some stromal cells that were adjacent to vessels (A, blue arrows). In primary prostate xenografts from mice injected with the PSA adenoviral construct, no immunostaining for luciferase was observed in endothelial cells (B, black arrows). Immunostaining for GFP was observed in the majority of endothelial cells in primary xenografts (C, black arrows, positive control). GR-CD31 (D–G) and PR-CD31 (H–J) colocalization was analyzed in primary prostate xenografts using the mouse/rabbit EnVision G/2 Double Stain System. GR expression was not detected in any prostate endothelial cells (E–G, black arrows). However, GR expression was observed in the nuclei of some stromal cells (E–G, blue arrows). PR expression was not detected in HPECs (I and J, black arrows). Specificity of the PR antibody was verified in breast cancer tissue samples (K, blue arrows). Bars, 10 μm.

The MMTV promoter is promiscuous, and MMTV-driven transcription can be stimulated by steroid nuclear receptors others than AR (i.e. GR and PR) (28,29,30). Consequently, expression of GR and PR nuclear receptors was analyzed by immunocytochemistry in primary xenografts of human benign prostate tissue. Expression of GR (Fig. 2, E–G, black arrows) or PR (Fig. 2, I and J, black arrows) was not observed in HPECs. As expected, GR expression was found in the nuclei of some stromal cells in the primary xenografts of human prostate (Fig. 2, E–G, blue arrows), which validated the specificity of the antibody. Human breast cancer tissue samples were used as a positive control for the anti-PR antibody, and PR immunostaining was apparent in the majority of nuclei of breast cancer cells (Fig. 2K, blue arrows).

These results, together with the high level of circulating testosterone (∼4 ng/ml) maintained in the host animals throughout the study, strongly suggest that the reporter construct activity was due to in vivo expression of functional AR in HPECs, consistent with the immunohistochemical analyses. However, the heterogeneous expression of AR may suggest the presence of functionally different subpopulations of endothelial cells in human prostate.

Isolation and characterization of HPECs in vitro

Primary cultures of HPECs were established from fresh surgical specimens using a Dynabead-based methodology (23). HPECs in primary culture showed endothelial cell morphology, functionality, and marker expression profiles comparable to HUVEC. Initially, HPECs in primary culture demonstrated an elongated appearance, which after 1 wk in culture changed to a more typical cobblestone morphology that exhibited a flattened appearance as the cells became confluent (Fig. 3A). HPECs in primary culture plated on a layer of Matrigel aligned with one another and, within 12–24 h, formed an anastomosing network of tubular structures that resembled a capillary plexus (Fig. 3B).

Figure 3.

Figure 3

Open in a new tab

Characterization of primary cultures of endothelial cells isolated from human benign prostate tissue. A, Morphology of HPECs after immunoselection and 1-wk culture. B, Endothelial tube network formed by primary cultures of HPECs cultured on Matrigel (120 μl/cm2) for 12–24 h. C, RT-PCR of RNA from primary cultures of HPECs, HUVECs, or LNCaP cells. PCR was performed using primers specific for human CD31, CD34, von Willebrand factor (vWF), ICAM, VEGFR1, VEGFR2, and PSA. GAPDH was used as a loading control. D, Immunostaining of HPEC and HUVECs, and benign prostate tissue sections, using antibodies specific for human CD31, CD34, and von Willebrand factor. Immunostaining in the absence of primary antibody provided negative controls (D, control). Black arrows indicate positively and negatively stained blood vessels in prostate tissue. Bars, 10 μm.

Primary cultures of HPECs were analyzed for expression of markers characteristic of human endothelial cells using RT-PCR and immunocytochemical analyses (Fig. 3, C and D). RT-PCR analyses demonstrated that cultured HPECs expressed mRNA encoding CD31, CD34, von Willebrand factor, intercellular adhesion molecule (ICAM), VEGF receptor (VEGFR) 1, and VEGFR2, but not PSA, which confirmed the absence of secretory epithelial cells in the cultures. A similar expression profile was observed in HUVECs. LNCaP cells were used as a negative control for expression of endothelial cell markers (Fig. 3C). Immunocytochemical analyses using specific antibodies against CD31, CD34, and von Willebrand factor in primary culture of HPECs and HUVECs confirmed expression of the endothelial cell-specific marker proteins (Fig. 3D). CD31 and CD34 immunostaining was localized to the cytoplasmic membrane in cultured HPECs and HUVECs, whereas von Willebrand factor immunostaining showed a characteristic granulated or punctate cytoplasmic pattern of localization, consistent with specific inclusion in Weibel-Palade bodies, a characteristic of endothelial cells (31). Specificity of these antibodies was confirmed by analysis of expression of CD31, CD34, and von Willebrand factor in formalin-fixed, paraffin-embedded tissue sections of benign human prostate (Fig. 3D). Immunostaining in the absence of primary antibody provided negative controls. Together, these results indicated that primary cultures of HPECs established from clinical specimens of human benign prostate contained predominantly endothelial cells.

HPECs express functional AR in vitro

AR expression in primary culture of HPECs was analyzed using RT-PCR, cDNA sequencing, immunocytochemistry, and Western blot analyses. RT-PCR experiments were performed using cDNA prepared from total RNA obtained from primary cultures of HPECs and specific primers designed for human AR. A single PCR amplification product of the size expected for AR (194 bp, position in the cDNA sequence: +3687 to +3881) was observed in primary cultures of HPECs (Fig. 4A). The specificity of the amplification reaction was verified using cDNA obtained from LNCaP cells that express mRNA for human AR (Fig. 4A, LNCaP). GAPDH was used as an internal control for the PCR analyses. Sequence analysis of the amplified fragment obtained from primary culture of HPECs demonstrated 100% sequence identity to the consensus human AR sequence (PubMed database, accession locus: NM_00044) (data not shown). Expression of AR at the protein level, and translocation of AR protein to the nucleus after DHT treatment, was demonstrated using immunocytochemistry in primary cultures of HPECs exposed for 24 h to vehicle (ethanol) or 5 nm DHT (Fig. 4B). In the absence of DHT, AR protein was localized primarily in the cytoplasm (Fig. 4B, i), whereas, in the presence of DHT, AR was localized to the nucleus (Fig. 4B, ii). As described for HPECs in human prostate tissue (Fig. 1), AR immunostaining was heterogeneous within primary cultures of HPECs; the majority of endothelial cell nuclei were positive for AR immunostaining (Fig. 4B, ii and iii), however, some did not contain immunodetectable AR (Fig. 4B, ii, black arrows). Colocalization of AR (red, Cy3) and von Willebrand factor (green, Cy2) confirmed the presence of AR protein in cultured HPECs (Fig. 4B, iv). In contrast, essentially all LNCaP cell nuclei immunostained for AR in the absence of DHT (Fig. 4B, v), and the intensity of immunostaining increased in the presence of DHT (Fig. 4B, vi and vii). Western blots analyses using nuclear extracts obtained from primary cultures of HPECs, and cultures of LNCaP cells, treated with 5 nm DHT demonstrated a single 110-kDa band for AR in both cell types (Fig. 4C). To establish a comparison between the levels of nuclear AR expression observed in the two cell types, a densitometric quantitation of the AR band of Western blots was performed. The results indicated that the level of AR expression in primary cultures of HPECs was approximately 17% the level of AR in LNCaP cells. Together, these results demonstrated that primary cultures of HPECs expressed AR in vitro, and AR translocated to the nucleus of prostate endothelial cells in the presence of DHT stimulation.

Figure 4.

Figure 4

Open in a new tab

AR expression in primary cultures of HPECs. A, RT-PCR of RNA from primary cultures of HPECs or LNCaP cells. PCR was performed using primers specific for human AR. B, Immunostaining of AR in HPECs and LNCaP cells. Cell cultures were incubated for 24 h with vehicle (-DHT, i and v) or 5 nm DHT (+DHT, ii and vi) before immunostaining. Black arrows indicate endothelial cells that did not immunostain for AR (B, ii, +DHT). Higher magnification images demonstrate clear nuclear immunostaining for AR in HPEC (B, iii) and LNCaP cells (B, vii). Colocalization of AR (red, Cy3) and von Willebrand factor (green, Cy2) confirmed the presence of AR in prostate endothelial cells (B, iv). Bars, 10 μm. C, Immunoblotting of nuclear extracts from HPECs or LNCaP cells after 24-h treatment with 5 nm DHT. Blots were reacted with anti-AR (AR). Topoisomerase I (Topo I) was used as a loading control for nuclear extracts.

Ligand-binding affinity of AR was analyzed in primary cultures of HPECs and LNCaP cells using the radiolabeled synthetic agonist/ligand, R1881. Levels of specific binding of R1881 to AR were estimated by subtracting background nonspecific binding from total binding. Specific R1881 binding to AR in cultured HPECs and LNCaP cells increased in a dose-dependent manner, and binding was saturated above ligand concentrations of 2 nm in both cell types (Fig. 5, A and C). Scatchard transformation of the binding data (specific R1881 binding/free ligand concentration vs. specific R1881 binding) resulted in a straight-line plot, indicative of a single ligand-binding site for R1881. Binding of R1881 had a Kd of 0.25 nm and 1.8 × 107 receptors per μg protein for cultured HPECs (Fig. 5B), and a Kd of 0.1 nm and 8.6 × 107 binding sites per μg protein for LNCaP cells (Fig. 5D). Consistent with the Western blot assay based estimate of the relative amount of receptor in HPECs compared with LNCaP cells, Scatchard analysis demonstrated that HPECs expressed 15–22% the number of binding sites per μg protein (1.8 × 107 binding sites per μg protein) observed in LNCaP cells (8.6 × 107 binding sites per μg protein) (Fig. 5E).

Figure 5.

Figure 5

Open in a new tab

Ligand-binding assay and functional characterization of AR in primary cultures of HPECs. Ligand-binding assays were performed in primary cultures of HPECs, and LNCaP cells using the radiolabeled synthetic agonist R1881. Specific binding of R1881 was estimated by subtraction of the nonspecific binding from the total binding of R1881. A, Specific [3H]R1881 binding determined by incubation of HPECs with R1881 at concentrations ranging from 0.01–6.0 nm. B, Scatchard plot of the specific binding data from graph A indicated a single binding site for R1881 to AR in primary cultures of HPECs with a kd of 0.25 nm. C, Specific [3H]R1881 binding determined by incubation of LNCaP cells with R1881 at concentrations ranging from 0.01–6.0 nm. D, Scatchard plot of the specific binding data from graph C indicated a single binding site for R1881 to AR in LNCaP cells with a kd of 0.1 nm. E, Number of binding sites of AR determined in HPECs and LNCaP cells. F, Relative luciferase reporter activities driven by MMTV or PSA promoters were determined in primary cultures of HPECs, LNCaP, and CV-1 cells in the absence (□, basal activity) and presence (▪, DHT inducible) of 5 nm DHT. Basal transcriptional activities (□) in HPECs, LNCaP, and CV-1 cells were adjusted to one. DHT-inducible transcriptional activities (▪) were presented in terms of fold induction (numbers over the black bars). G, Inhibition of the DHT-inducible luciferase reporter activity driven by the MMTV promoter by increasing concentrations of flutamide (0.2, 2, 20, and 200 μm) in primary culture of HPECs. Relative luciferase activity is presented as percentage of the control. Statistical evaluation of the data were performed using the ANOVA test. *, P ≤ 0.05; **, P ≤ 0.01 (n = 3). RLU, Relative luciferase units.

Functional activity of AR in primary cultures of HPECs and LNCaP cells was analyzed by measurement of AR-mediated transcriptional activity using an adenoviral MMTV- or PSA-promoter driven luciferase gene reporter construct (Fig. 5F) (32). A steroid receptor-deficient monkey kidney cell line, CV-1 cells, was used as a negative control for AR-mediated transcriptional activity studies (33,34,35). DHT increased the reporter activity of the MMTV-promoter driven construct by 29-fold in primary cultures of HPECs (Fig. 5F, HPEC, MMTV) and 58-fold in LNCaP cells (Fig. 5F, LNCaP, MMTV). In contrast, DHT did not stimulate expression of the luciferase reporter driven by the epithelial cell-specific PSA promoter in primary cultures of HPECs (Fig. 5F, HPEC, PSA). However, DHT increased PSA promoter-driven reporter activity 20-fold in LNCaP cells (Fig. 5F, LNCaP, PSA), which indicated cell type specific AR-mediated transactivation in different cell types isolated from human prostate. As expected, DHT did not stimulate expression of the luciferase reporter driven by the MMTV or PSA promoter in CV-1 cells (Fig 5F, CV-1). Preincubation of primary cultures of prostate endothelial cells with the antiandrogen flutamide inhibited the MMTV-driven luciferase reporter activity in a dose-dependent manner (Fig. 5G), indicating that the transcriptional activity of the MMTV-driven luciferase gene reporter was AR mediated, and not due to GR or PR.

Androgen, through AR, increased proliferation in primary culture of HPECs

To determine the range of biological properties modulated by androgen in primary cultures of HPECs, assays of proliferation (Ki-67 expression measurement) and endothelial cell tube formation in Matrigel were performed in the presence of increasing concentrations of DHT (Fig. 6). DHT increased significantly the proliferation index of primary cultures of HPECs in a dose-dependent manner (Fig. 6A). Proliferation indices reached a plateau at 1.0 nm DHT, with no further increase in proliferation observed for higher concentrations of DHT (10 nm). Moreover, the increase in proliferation index induced by stimulation by 1 nm DHT was inhibited by the antiandrogen flutamide in a dose-dependent manner (Fig. 6B). Interestingly, DHT exhibited no effect on the ability of HPECs to form tubular structures in Matrigel (Fig. 6, C and D). Consequently, androgens, through AR, modulate only a subset of biological properties of primary culture of HPECs.

Figure 6.

Figure 6

Open in a new tab

Effect of DHT on proliferation and endothelial tube formation in Matrigel in primary cultures of HPECs. A, The effect of androgen on proliferation of primary cultures of HPECs was analyzed by quantifying the percentage of Ki-67-expressing cells in 10 different microscopical fields of cells exposed to increasing concentrations of DHT (0, 0.01, 0.1, 1, and 10 nm). B, Inhibition of DHT-stimulated (1 nm) proliferation of HPECs by flutamide (20 and 200 μm). Ki-67-expressing cells were quantified in 10 different microscopical fields and presented as percentage of the control (presence of 1 nm DHT and absence of flutamide). C, Effect of androgen on endothelial tube formation in Matrigel by primary cultures of HPECs was analyzed in the presence of increasing concentrations of DHT (0, 0.01, 0.1, 1, and 10 nm). D, Quantification of relative tube length formed by primary cultures of HPECs exposed to increasing concentration of DHT. Statistical evaluation of the data was performed using the ANOVA test. *, P ≤ 0.05; **, P ≤ 0.01 (n = 4).

Discussion

Expression and functionality of AR in human prostate tissue have been described in detail for the epithelial and stromal compartments (32,36,37,38,39,40). The response of prostate tissue to androgen deprivation is characterized, in general, by remarkable changes in gene expression in prostate epithelial cells that correlated with apoptosis (41). However, these castration-induced effects on prostate epithelial cells might be, at least in part, an indirect response to castration-induced perturbation of the vasculature of the prostate gland that precedes the onset of epithelial cell apoptosis by several days (7,8,11,41). Mechanistic characterization of the androgen deprivation-induced reduction in vascular function has been hampered by a lack of identification of the specific sensitivities of the individual AR-expressing cellular compartments in the prostate. A paracrine mechanism for endothelial cell death in response to androgen deprivation was anticipated in light of the report of Prins et al. (10) in 1991 that AR was not expressed in prostate endothelial cells in the adult rat. However, El-Alfy et al. (20) reported in 1999 that AR was present in HPECs, which suggested that endothelial cell apoptosis could be, at least in part, a direct response to androgen deprivation.

This report provides an in-depth characterization of the in vivo and in vitro expression and functionality of AR in HPECs. Initially, expression of AR protein in human benign prostate and CaP tissue samples obtained from clinical specimens was analyzed using immunohistochemistry. Consistent with the previous report (20), AR expression was heterogeneous within HPECs, suggesting that the endothelial cell compartment may exhibit several functional phenotypes. Multiple populations of endothelial cells could have different roles and responses in modulating androgen action in the vasculature of human prostate. Although vascular endothelial cells share common functions, considerable heterogeneity exists both structurally and functionally along the length of the vascular tree. Endothelial cells form tight, continuous monolayers in the brain, and discontinuous layers with intercellular gaps or fenestrae in kidney and spleen (42,43,44). There also are clear differences between endothelial cells in the microvasculature and macrovasculature within the vascular beds of the same organ (45). In vivo studies have demonstrated heterogeneous phenotypes in lung endothelial cells along the pulmonary vascular tree that reflect functional adaptation to local requirements and environmental factors. However, the heterogeneous expression of AR in HPECs appears unrelated to localization of the vasculature within the prostate because differential AR expression in endothelial cells was observed within single vessels. AR expression also appears similar in the microvasculature and macrovasculature of the prostate because AR positive endothelial cells were found in both the microvasculature and macrovasculature of clinical specimens of human benign prostate and CaP. Therefore, one possible explanation for the heterogeneous expression of AR in human prostate vasculature would be a cell cycle stage-specific expression of AR in HPECs. However, more studies are necessary to address possible mechanisms responsible for differential expression of AR in HPECs.

In vivo functionality of AR in HPECs was demonstrated using adenoviral constructs of MMTV- or PSA-driven luciferase gene reporters injected via the tail vein into SCID mice bearing human prostate xenografts (21,22). AR-mediated expression of the luciferase gene was observed in endothelial cells of human prostate xenografts in mice injected with the MMTV-driven luciferase adenoviral construct, but not the PSA-driven adenoviral construct, confirming a cell type-specific AR-mediated transactivation in prostate endothelial cells. MMTV promoter activity can be stimulated by steroid nuclear receptors other than AR, such as GR and PR (28,29,30), therefore, immunocytochemical analyses of expression of the nuclear receptors GR and PR were performed in primary xenografts of benign human prostate tissue. GR and PR were not expressed in HPECs. Furthermore, the marked nuclear localization of AR observed in endothelial cells in the presence of the high levels of circulating testosterone in the host animals strongly suggests that AR is responsible for the in vivo transcription of the MMTV-driven reporter. In addition, the high level of expression of the reporter construct suggests the usefulness of this delivery system for further studies of the endothelium in vivo and possible usefulness of therapeutic modalities directed to the endothelium.

Expression of AR in HPECs was analyzed in vitro in primary cultures established from fresh surgical specimens of human benign prostate tissue. Consistent with the in vivo data, primary cultures of HPECs showed heterogeneous expression of AR, with AR expressed at approximately 15–22% of the level observed in LNCaP cells as determined by quantitation of Western blots and by measuring the number of receptors using hormone binding. Moreover, in vivo immunostaining analyses indicated that AR was expressed at a comparable level of intensity in HPECs and human prostate stromal cells.

Ligand-binding assays confirmed the presence of a single high-affinity binding site for R1881 in the AR of HPECs with a Kd of 0.2 nm, a value in the range of Kd for wild-type AR previously described in other cellular models (46). Ligand-induced translocation of AR from the cytoplasm to the nucleus is a critical step in the pathway of androgen-mediated activation/repression of gene transcription (47). Ligand-induced nuclear translocation of AR was observed in primary culture of HPECs in response to incubation with DHT. Finally, AR-mediated transcriptional activation was demonstrated by infecting primary cultures of HPECs with an adenoviral MMTV promoter-driven luciferase reporter gene. These results were consistent with the in vivo transactivation data and confirmed the maintenance of functional activity of AR in cultured HPECs. Interestingly, the 4- to 6-fold difference in the number of receptors between LNCaP cells and primary cultures of HPECs was associated with only a 2-fold higher level of AR-mediated transcriptional activation in LNCaP cells compared with endothelial cells. This observation further supports the hypothesis of very active AR signaling in HPECs.

DHT significantly increased proliferation of HPECs in primary culture. Previous studies in the adult rat ventral prostate suggested that castration reduced endothelial cell number, and the endothelial cell proliferation rate, and that both endpoints were normalized by testosterone treatment (48,49). Similarly, castration decreased, and testosterone treatment rapidly normalized, blood flow to the adult rat ventral prostate (7,8). These studies suggested that the vasculature was regulated, directly or indirectly, by androgens. However, in rat ventral prostate, AR expression was reported only in epithelial and stromal cells, but not in blood vessels (10). Therefore, the vascular effects appeared secondary to changes in other prostate cell compartments (7,8,11,12). Our results demonstrate that HPECs express functional AR in vivo and in vitro, and that androgen modulates in vitro endothelial cell proliferation and gene expression in a cell type-specific manner. Demonstration that the antiandrogen flutamide inhibits in vitro proliferation of prostate endothelial cells in culture supports the hypothesis that androgens directly modulate HPEC proliferation through activation of AR within the endothelial cell. However, our studies cannot eliminate the possibility of an indirect contribution of androgens to maintaining endothelial cell homeostasis in vivo. In support of this hypothesis, castration-induced involution of tumor vessels of androgen-dependent Shionogi carcinoma was concomitant with decreased VEGF expression in tumor epithelial cells (12). Therefore, further in vivo analyses will be required to determine the relative contributions of both mechanisms, a direct endothelial cell AR-mediated effect of androgen on prostate vasculature, and an indirect effect of androgen mediated through another AR-expressing prostate cell.

Angiogenesis is a complex process that requires not only endothelial cell proliferation, but also sprouting, migration, and differentiation of endothelial cells into tube structures with production of a basement membrane matrix around the vessels (50,51,52,53,54). This study demonstrated that androgen has a differential effect on the processes involved in “angiogenesis” of HPECs in primary culture because androgen increased endothelial cell proliferation but did not affect endothelial tube formation. However, elucidation of the role of androgen signaling in the specific steps in angiogenesis in vitro and in vivo will require further studies.

Considerable evidence has been reported to suggest that tumor mass is under strict control of the microvascular endothelium (11,55), and that recruitment and invasion of capillary sized vessels stimulated by signals from tumor cells are reinforced by potent paracrine effects of the endothelial cells themselves (56). The transition from androgen-stimulated to castration-recurrent CaP is associated with increased microvessel density (55,57). Nascent endothelial cells produce and release into their extracellular matrix, as well as into their tumor microenvironment, a variety of potent growth and survival factors (56,58). At least 20 endothelial cell-derived paracrine factors have been described (23,58), including basic fibroblast growth factor, platelet-derived growth factor, IGF-I, heparin-binding epithelial growth factor, and IL-6. Therefore, understanding the cellular and molecular mechanisms of AR-mediated androgen action, and the modulation of gene expression by androgen, in HPECs isolated from benign and malignant tissue specimens could contribute to a better understanding of the biological role of the HPECs in normal and pathological conditions.

In summary, our results present the first evidence of expression of functional AR in HPECs in vivo and in vitro. Moreover, these studies indicate an important role of androgen in modulating homeostasis of HPECs in vitro and suggest that the disruption of the unique signaling mechanism in the vasculature of the human prostate may provide a primary therapeutic target for ADT.

Acknowledgments

We thank Dr. James L. Mohler for the critical review of the manuscript and Suzanne Cook for technical assistance.

Footnotes

This work was supported by PO1-CA77739 (to G.J.S.).

Disclosure Statement: A.G., A.W., P.S., V.P.M., W.J.H., and S.A.O. have nothing to declare. G.J.S. has a financial interest in AndroBioSys, Inc.

First Published Online February 21, 2008

Abbreviations: ADT, Androgen deprivation therapy; AR, androgen receptor; CaP, prostate cancer; CMV, cytomegalovirus; DHT, dihydrotestosterone; FBS, fetal bovine serum; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GFP, green fluorescence protein; GR, glucocorticoid receptor; HPEC, human prostate endothelial cell; HRP, horseradish peroxidase; HUVEC, human umbilical vein endothelial cell; ICAM, intercellular adhesion molecule; ifu, infectious units; Kd, dissociation constant; MMTV, mouse mammary tumor virus; PR, progesterone receptor; PSA, prostate-specific antigen; RPCI, Roswell Park Cancer Institute; SCID, severe combined immunodeficient; TBS, Tris-buffered saline; VEGF, vascular endothelial growth factor; VEGFR, vascular endothelial growth factor receptor.

References

  1. Huggins C, Hodges CV 2002 Studies on prostatic cancer: I. The effect of castration, of estrogen and of androgen injection on serum phosphatases in metastatic carcinoma of the prostate. 1941. J Urol 168:9–12 [DOI] [PubMed] [Google Scholar]
  2. Crawford ED 1989 Combined androgen blockade. Urology 34:22–26 [DOI] [PubMed] [Google Scholar]
  3. Prins GS, Jung MH, Vellanoweth RL, Chatterjee B, Roy AK 1996 Age-dependent expression of the androgen receptor gene in the prostate and its implication in glandular differentiation and hyperplasia. Dev Genet 18:99–106 [DOI] [PubMed] [Google Scholar]
  4. Prins GS, Birch L 1995 The developmental pattern of androgen receptor expression in rat prostate lobes is altered after neonatal exposure to estrogen. Endocrinology 136:1303–1314 [DOI] [PubMed] [Google Scholar]
  5. Thompson TC, Cunha GR, Shannon JM, Chung LW 1986 Androgen-induced biochemical responses in epithelium lacking androgen receptors: characterization of androgen receptors in the mesenchymal derivative of urogenital sinus. J Steroid Biochem 25:627–634 [DOI] [PubMed] [Google Scholar]
  6. Feldman BJ, Feldman D 2001 The development of androgen-independent prostate cancer. Nat Rev Cancer 1:34–45 [DOI] [PubMed] [Google Scholar]
  7. Shabsigh A, Chang DT, Heitjan DF, Kiss A, Olsson CA, Puchner PJ, Buttyan R 1998 Rapid reduction in blood flow to the rat ventral prostate gland after castration: preliminary evidence that androgens influence prostate size by regulating blood flow to the prostate gland and prostatic endothelial cell survival. Prostate 36:201–206 [DOI] [PubMed] [Google Scholar]
  8. Lekas E, Johansson M, Widmark A, Bergh A, Damber JE 1997 Decrement of blood flow precedes the involution of the ventral prostate in the rat after castration. Urol Res 25:309–314 [DOI] [PubMed] [Google Scholar]
  9. Shabisgh A, Tanji N, D'Agati V, Burchardt M, Rubin M, Goluboff ET, Heitjan D, Kiss A, Buttyan R 1999 Early effects of castration on the vascular system of the rat ventral prostate gland. Endocrinology 140:1920–1926 [DOI] [PubMed] [Google Scholar]
  10. Prins GS, Birch L, Greene GL 1991 Androgen receptor localization in different cell types of the adult rat prostate. Endocrinology 129:3187–3199 [DOI] [PubMed] [Google Scholar]
  11. Folkman J 1998 Is tissue mass regulated by vascular endothelial cells? Prostate as the first evidence. Endocrinology 139:441–442 [DOI] [PubMed] [Google Scholar]
  12. Jain RK, Safabakhsh N, Sckell A, Chen Y, Jiang P, Benjamin L, Yuan F, Keshet E 1998 Endothelial cell death, angiogenesis, and microvascular function after castration in an androgen-dependent tumor: role of vascular endothelial growth factor. Proc Natl Acad Sci USA 95:10820–10825 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Liang T, Hoyer S, Yu R, Soltani K, Lorincz AL, Hiipakka RA, Liao S 1993 Immunocytochemical localization of androgen receptors in human skin using monoclonal antibodies against the androgen receptor. J Invest Dermatol 100:663–666 [DOI] [PubMed] [Google Scholar]
  14. Blauer M, Vaalasti A, Pauli SL, Ylikomi T, Joensuu T, Tuohimaa P 1991 Location of androgen receptor in human skin. J Invest Dermatol 97:264–268 [DOI] [PubMed] [Google Scholar]
  15. Laine M, Blauer M, Ylikomi T, Tuohimaa P, Aitasalo K, Happonen RP, Tenovuo J 1993 Immunohistochemical demonstration of androgen receptors in human salivary glands. Arch Oral Biol 38:299–302 [DOI] [PubMed] [Google Scholar]
  16. Abu EO, Horner A, Kusec V, Triffitt JT, Compston JE 1997 The localization of androgen receptors in human bone. J Clin Endocrinol Metab 82:3493–3497 [DOI] [PubMed] [Google Scholar]
  17. Mantalaris A, Panoskaltsis N, Sakai Y, Bourne P, Chang C, Messing EM, Wu JH 2001 Localization of androgen receptor expression in human bone marrow. J Pathol 193:361–366 [DOI] [PubMed] [Google Scholar]
  18. Schultheiss D, Badalyan R, Pilatz A, Gabouev AI, Schlote N, Wefer J, von Wasielewski R, Mertsching H, Sohn M, Stief CG, Jonas U 2003 Androgen and estrogen receptors in the human corpus cavernosum penis: immunohistochemical and cell culture results. World J Urol 21:320–324 [DOI] [PubMed] [Google Scholar]
  19. Sinha-Hikim I, Taylor WE, Gonzalez-Cadavid NF, Zheng W, Bhasin S 2004 Androgen receptor in human skeletal muscle and cultured muscle satellite cells: up-regulation by androgen treatment. J Clin Endocrinol Metab 89:5245–5255 [DOI] [PubMed] [Google Scholar]
  20. El-Alfy M, Luu-The V, Huang XF, Berger L, Labrie F, Pelletier G 1999 Localization of type 5 17β-hydroxysteroid dehydrogenase, 3β-hydroxysteroid dehydrogenase, and androgen receptor in the human prostate by in situ hybridization and immunocytochemistry. Endocrinology 140:1481–1491 [DOI] [PubMed] [Google Scholar]
  21. Gray DR, Huss WJ, Yau JM, Durham LE, Werdin ES, Funkhouser Jr WK, Smith GJ 2004 Short-term human prostate primary xenografts: an in vivo model of human prostate cancer vasculature and angiogenesis. Cancer Res 64:1712–1721 [DOI] [PubMed] [Google Scholar]
  22. Presnell SC, Werdin ES, Maygarden S, Mohler JL, Smith GJ 2001 Establishment of short-term primary human prostate xenografts for the study of prostate biology and cancer. Am J Pathol 159:855–860 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Stachon A, Schluter T, Koller M, Weisser H, Krieg M 2001 Primary culture of microvascular endothelial cells from human benign prostatic hyperplasia. Prostate 48:156–164 [DOI] [PubMed] [Google Scholar]
  24. Godoy A, Ormazabal V, Moraga-Cid G, Zuniga FA, Sotomayor P, Barra V, Vasquez O, Montecinos V, Mardones L, Guzman C, Villagran M, Aguayo LG, Onate SA, Reyes AM, Carcamo JG, Rivas CI, Vera JC 2007 Mechanistic insights and functional determinants of the transport cycle of the ascorbic acid transporter SVCT2. Activation by sodium and absolute dependence on bivalent cations. J Biol Chem 282:615–624 [DOI] [PubMed] [Google Scholar]
  25. Godoy A, Ulloa V, Rodriguez F, Reinicke K, Yanez AJ, Garcia Mde L, Medina RA, Carrasco M, Barberis S, Castro T, Martinez F, Koch X, Vera JC, Poblete MT, Figueroa CD, Peruzzo B, Perez F, Nualart F 2006 Differential subcellular distribution of glucose transporters GLUT1–6 and GLUT9 in human cancer: ultrastructural localization of GLUT1 and GLUT5 in breast tumor tissues. J Cell Physiol 207:614–627 [DOI] [PubMed] [Google Scholar]
  26. Nualart F, Godoy A, Reinicke K 1999 Expression of the hexose transporters GLUT1 and GLUT2 during the early development of the human brain. Brain Res 824:97–104 [DOI] [PubMed] [Google Scholar]
  27. Brinkmann AO, Bolt J, van Steenbrugge GJ, Kuiper GG, de Boer W, Mulder E 1987 Characterization of androgen receptors in a transplantable human prostatic adenocarcinoma (PC-82). Prostate 10:133–143 [DOI] [PubMed] [Google Scholar]
  28. von der Ahe D, Janich S, Scheidereit C, Renkawitz R, Schutz G, Beato M 1985 Glucocorticoid and progesterone receptors bind to the same sites in two hormonally regulated promoters. Nature 313:706–709 [DOI] [PubMed] [Google Scholar]
  29. Otten AD, Sanders MM, McKnight GS 1988 The MMTV LTR promoter is induced by progesterone and dihydrotestosterone but not by estrogen. Mol Endocrinol 2:143–147 [DOI] [PubMed] [Google Scholar]
  30. Cato AC, Miksicek R, Schutz G, Arnemann J, Beato M 1986 The hormone regulatory element of mouse mammary tumour virus mediates progesterone induction. EMBO J 5:2237–2240 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Wagner DD, Olmsted JB, Marder VJ 1982 Immunolocalization of von Willebrand protein in Weibel-Palade bodies of human endothelial cells. J Cell Biol 95:355–360 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Cano P, Godoy A, Escamilla R, Dhir R, Onate SA 2007 Stromal-epithelial cell interactions and androgen receptor-coregulator recruitment is altered in the tissue microenvironment of prostate cancer. Cancer Res 67:511–519 [DOI] [PubMed] [Google Scholar]
  33. Fuhrmann U, Bengtson C, Repenthin G, Schillinger E 1992 Stable transfection of androgen receptor and MMTV-CAT into mammalian cells: inhibition of cat expression by anti-androgens. J Steroid Biochem Mol Biol 42:787–793 [DOI] [PubMed] [Google Scholar]
  34. Rundlett SE, Wu XP, Miesfeld RL 1990 Functional characterizations of the androgen receptor confirm that the molecular basis of androgen action is transcriptional regulation. Mol Endocrinol 4:708–714 [DOI] [PubMed] [Google Scholar]
  35. Warriar N, Page N, Koutsilieris M, Govindan MV 1993 Interaction of antiandrogen-androgen receptor complexes with DNA and transcription activation. J Steroid Biochem Mol Biol 46:699–711 [DOI] [PubMed] [Google Scholar]
  36. Mohler JL, Chen Y, Hamil K, Hall SH, Cidlowski JA, Wilson EM, French FS, Sar M 1996 Androgen and glucocorticoid receptors in the stroma and epithelium of prostatic hyperplasia and carcinoma. Clin Cancer Res 2:889–895 [PubMed] [Google Scholar]
  37. Gerdes MJ, Dang TD, Lu B, Larsen M, McBride L, Rowley DR 1996 Androgen-regulated proliferation and gene transcription in a prostate smooth muscle cell line (PS-1). Endocrinology 137:864–872 [DOI] [PubMed] [Google Scholar]
  38. Sar M, Lubahn DB, French FS, Wilson EM 1990 Immunohistochemical localization of the androgen receptor in rat and human tissues. Endocrinology 127:3180–3186 [DOI] [PubMed] [Google Scholar]
  39. Chodak GW, Kranc DM, Puy LA, Takeda H, Johnson K, Chang C 1992 Nuclear localization of androgen receptor in heterogeneous samples of normal, hyperplastic and neoplastic human prostate. J Urol 147:798–803 [DOI] [PubMed] [Google Scholar]
  40. Brolin J, Skoog L, Ekman P 1992 Immunohistochemistry and biochemistry in detection of androgen, progesterone, and estrogen receptors in benign and malignant human prostatic tissue. Prostate 20:281–295 [DOI] [PubMed] [Google Scholar]
  41. Buttyan R, Shabsigh A, Perlman H, Colombel M 1999 Regulation of apoptosis in the prostate gland by androgenic steroids. Trends Endocrinol Metab 10:47–54 [DOI] [PubMed] [Google Scholar]
  42. Risau W 1995 Differentiation of endothelium. FASEB J 9:926–933 [PubMed] [Google Scholar]
  43. Garlanda C, Dejana E 1997 Heterogeneity of endothelial cells. Specific markers. Arterioscler Thromb Vasc Biol 17:1193–1202 [DOI] [PubMed] [Google Scholar]
  44. Cines DB, Pollak ES, Buck CA, Loscalzo J, Zimmerman GA, McEver RP, Pober JS, Wick TM, Konkle BA, Schwartz BS, Barnathan ES, McCrae KR, Hug BA, Schmidt AM, Stern DM 1998 Endothelial cells in physiology and in the pathophysiology of vascular disorders. Blood 91:3527–3561 [PubMed] [Google Scholar]
  45. Hewett PW, Murray JC 1993 Human microvessel endothelial cells: isolation, culture and characterization. In Vitro Cell Dev Biol Anim 29A:823–830 [DOI] [PubMed] [Google Scholar]
  46. Tan J, Sharief Y, Hamil KG, Gregory CW, Zang DY, Sar M, Gumerlock PH, DeVere White RW, Pretlow TG, Harris SE, Wilson EM, Mohler JL, French FS 1997 Dehydroepiandrosterone activates mutant androgen receptors expressed in the androgen-dependent human prostate cancer xenograft CWR22 and LNCaP cells. Mol Endocrinol 11:450–459 [DOI] [PubMed] [Google Scholar]
  47. Kemppainen JA, Lane MV, Sar M, Wilson EM 1992 Androgen receptor phosphorylation, turnover, nuclear transport, and transcriptional activation. Specificity for steroids and antihormones. J Biol Chem 267:968–974 [PubMed] [Google Scholar]
  48. English HF, Drago JR, Santen RJ 1985 Cellular response to androgen depletion and repletion in the rat ventral prostate: autoradiography and morphometric analysis. Prostate 7:41–51 [DOI] [PubMed] [Google Scholar]
  49. Franck-Lissbrant I, Haggstrom S, Damber JE, Bergh A 1998 Testosterone stimulates angiogenesis and vascular regrowth in the ventral prostate in castrated adult rats. Endocrinology 139:451–456 [DOI] [PubMed] [Google Scholar]
  50. Folkman J, Haudenschild C 1980 Angiogenesis in vitro. Nature 288:551–556 [DOI] [PubMed] [Google Scholar]
  51. Madri JA, Williams SK 1983 Capillary endothelial cell cultures: phenotypic modulation by matrix components. J Cell Biol 97:153–165 [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Kramer RH, Fuh GM, Bensch KG, Karasek MA 1985 Synthesis of extracellular matrix glycoproteins by cultured microvascular endothelial cells isolated from the dermis of neonatal and adult skin. J Cell Physiol 123:1–9 [DOI] [PubMed] [Google Scholar]
  53. Herbst TJ, McCarthy JB, Tsilibary EC, Furcht LT 1988 Differential effects of laminin, intact type IV collagen, and specific domains of type IV collagen on endothelial cell adhesion and migration. J Cell Biol 106:1365–1373 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Grant DS, Tashiro K, Segui-Real B, Yamada Y, Martin GR, Kleinman HK 1989 Two different laminin domains mediate the differentiation of human endothelial cells into capillary-like structures in vitro. Cell 58:933–943 [DOI] [PubMed] [Google Scholar]
  55. Lissbrant IF, Stattin P, Damber JE, Bergh A 1997 Vascular density is a predictor of cancer-specific survival in prostatic carcinoma. Prostate 33:38–45 [DOI] [PubMed] [Google Scholar]
  56. Rak J, Filmus J, Kerbel RS 1996 Reciprocal paracrine interactions between tumour cells and endothelial cells: the ‘angiogenesis progression’ hypothesis. Eur J Cancer 32A:2438–2450 [DOI] [PubMed] [Google Scholar]
  57. Gustavsson H, Welen K, Damber JE 2005 Transition of an androgen-dependent human prostate cancer cell line into an androgen-independent subline is associated with increased angiogenesis. Prostate 62:364–373 [DOI] [PubMed] [Google Scholar]
  58. Hamada J, Cavanaugh PG, Lotan O, Nicolson GL 1992 Separable growth and migration factors for large-cell lymphoma cells secreted by microvascular endothelial cells derived from target organs for metastasis. Br J Cancer 66:349–354 [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Endocrinology are provided here courtesy of The Endocrine Society

RESOURCES