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Original Studies |
Department of Clinical Physiopathology, Endocrinology Unit (C.C., R.S., M.S.) and Andrology Unit (M.M., M.L., G.F.); Department of Anatomy Histology and Forensic Medicine (G.B.V., M.G.), University of Florence, Florence; and Department of Clinical and Experimental Medicine (T.B.), University of Catanzaro, Catanzaro, Italy
Address correspondence and requests for reprints to: Mario Maggi, Department of Clinical Physiopathology, Andrology Unit, University of Florence, Viale Pieraccini, 6, 50139-Florence, Italy. E-mail: m.maggi{at}dfc.unifi.it
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Abstract |
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Introduction |
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Up to now, it is clear that androgens play a pivotal role in
controlling prostate growth and differentiation, not only during fetal
life and childhood but also in adult life. Hence, it has been proposed
that increased prostatic concentration of androgens, or increased
androgen responsiveness, causes BPH. However, different androgen
ablation strategies resulted in a modest decrease of the hyperplastic
prostate volume and were of limited effectiveness in reducing clinical
symptoms. In particular, although blockage of 5
-reductase type 2
activity by finasteride induced a consistent decrease in
the prostatic concentration of dihydrotestosterone (1, 2), it decreases
prostate volume by only 27% after 3 yr of therapy (3). During the last
few years, it became evident that both androgen-dependent and
androgen-independent growth factors promote prostate enlargement by
inducing cell proliferation or reducing apoptosis (4, 5).
Therefore, new therapeutic strategies, aimed at reducing intraprostatic
growth factor signaling, are needed for an efficient treatment of
BPH.
The activated form of vitamin D, vitamin D3, seems to be a promising candidate for BPH therapy. Indeed, it has emerged as one of the most potent growth regulatory molecules in prostate. Vitamin D3 binds to nuclear vitamin D receptor, present in both epithelial and stromal cells, and inhibits growth (6). The presence of nuclear vitamin D receptors has been showed in primary cultures of human prostatic cells (6), as well as in human prostate cancer lines (7). Epidemiological studies suggested a relationship between vitamin D3 deficiency or vitamin D3 receptor gene polymorphism and prostate cancer risk (8, 9), focusing the attention on a possible therapeutic or chemo-preventive use of vitamin D3 in cancer. However, vitamin D3 cannot be used as an anticancer drug, because of the fact that the drug causes hypercalcemia.
In the last few years, several vitamin D3 derivative molecules that have the same effect but are less hypercalcemic have been developed. Among these compounds, it has been demonstrated that 1,25-dihydroxy-16ene-23yne D3, named analogue (V), is 7-fold more potent than vitamin D3 in terms of differentiation and inhibition of clonal proliferation of HL60 cells, retaining only 2–3% of its calcemic activity (10). Therefore, analogue (V) has been proposed as a candidate drug for the treatment of promyelocytic acute leukemia (11, 12). Interestingly, the antiproliferative effect of analogue (V) has been demonstrated in prostate cancer cells in vivo (13).
The aim of the present study is to investigate the in vitro effects of vitamin D3 and its aforementioned analogue, analogue (V), on cell cultures derived from patients undergoing surgery for BPH. Effects on both cell proliferation and apoptosis have been studied in basal conditions and after treatment with the potent mitogen keratinocyte growth factor (KGF). KGF is a member of the fibroblast growth factor (FGF) family expressed by normal prostate (14, 15) and overexpressed by hormone insensitive prostate carcinomas (16). FGFs have been demonstrated to stimulate tyrosine phosphorylation of their own receptors, upon ligand binding (17).
We previously demonstrated not only the presence of specific transcripts for KGF and its receptor in human hyperplastic prostate tissue but also the potent mitogenic activity of KGF on BPH cells (15). Therefore, in the present study, we focused also on interactions among vitamin D3, analogue (V), and KGF, in terms of regulation of cell proliferation and cell death. In addition, the molecular mechanisms underlying these phenomena have been partially clarified.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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MEM, PBS, BSA, glutamine, antibiotics, collagenase type IV,
trypan blue, vitamin D3, reagents for
intracellular calcium measurement, electron microscopy, and
immunocytochemistry were from Sigma (St. Louis, MO). FBS
was obtained by Unipath (Bedford, England). Human KGF, recombinant,
(KGF), and BM-enhanced-chemiluminescence system were
purchased from Roche Molecular Biochemicals
Biochemica (Mannheim, Germany). Analogue 1,25-dihydroxy-16ene-23yne
D3 (V) was from La Roche Molecular Biochemicals (Indianapolis, IN). Mouse antihuman monoclonal
antibody against bcl-2, smooth-muscle actin, vimentin, and
cytokeratin; and rabbit antihuman polyclonal antibodies against desmin
and factor VIII were purchased from DAKO Corp.,
(Carpinteria, CA). Apop Tag kit was from Oncor (MD). Horse
radish peroxidase-conjugated monoclonal antiphosphotyrosine antibody
(PY20-HRP) was purchased from ICN Biomedicals, Inc.
(Costa Mesa, CA) and anti-KGF receptor (anti-KGF-R) antibody
from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).
Reagents for sodium dodecyl sulphate-polyacrylamide gel electrophoresis
(SDS-PAGE) and for protein measurement were from Bio-Rad Laboratories, Inc. (Hercules, CA). Fura-2/AM was obtained from
Calbiochem (La Jolla, CA). Plasticware for cell cultures
was purchased from Falcon (Oxnard, CA). Disposable filtration units for
growth media preparation were purchased from PBI International (Milan,
Italy).
Cell cultures and tissue
BPH cells were obtained from prostate tissues derived from five patients, who underwent suprapubic adenomectomy for BPH. Patients did not receive any pharmacological treatment in the 3 months preceding surgery. They were prepared as previously described (15). Briefly, tissues were cut in small fragments and treated overnight with 2 mg/mL bacterial collagenase. Fragments were then extensively washed in PBS and cultured in MEM supplemented with 10% heat-inactivated FBS, 2 mmol/L glutamine, 100 U/mL penicillin, and 100 µg/mL streptomycin in a fully humidified atmosphere of 95% air-5% CO2.
Cells began to emerge within 1 week and were used within the fifth
passage. Specific antibodies were used to characterize BPH cells. They
showed positive staining for smooth-muscle actin, vimentin, and
desmin, suggesting fibromuscular morphological features. Conversely,
they were negative for epithelial and endothelial markers such as
cytokeratin and factor VIII (data not shown).
Cell proliferation assay
For growth measurement, 6 x 104 cells were seeded onto 12-well plates in growth medium. After 24 h, the growth medium was removed, the cells were accurately washed in PBS and incubated in phenol red- and serum-free medium containing 0.1% BSA. After 24 h, increasing concentrations (1–100 ng/mL ) of KGF were added with or without a fixed concentration (10 nmol/L) of vitamin D3 or analogue (V). Experiments were also performed using increasing concentrations (0.1–100 nmol/L) of vitamin D3 or analogue (V) alone. Cells in phenol red- and serum-free medium containing 0.1% BSA were used as basal controls. After 48 h, cells were trypsinized, and each experimental point was derived from counting in the hemocytometer and then averaging at least five different fields for each well. In the same experiment, each experimental point was repeated in duplicate or triplicate. Experiments were repeated at least twice in more than one preparation of BPH cells. Cell growth results are expressed as percentage (±SE) of the growth of their relative controls.
SDS-PAGE and Western blot analysis
After the different treatments, cells were scraped, centrifuged, and resuspended in lysis buffer [20 mmol/L Tris (pH 7.4), 150 mmol/L NaCl, 0.25% Nonidet P40, 1 mmol/L Na3VO4, 1 mmol/L phenylmethylsulfonylfluoride]. After protein measurement, performed according to the manufacturer’s instruction (Bio-Rad Laboratories, Inc. kit), aliquots containing about 25 µg of proteins were diluted in reducing 2x SB [Laemmli’s sample buffer = 62.5 mmol/L Tris (pH 6.8), 10% glycerol, 20% SDS, 2.5% pyronin, and 200 mmol/L dithiothreitol], boiled, and loaded onto 8% polyacrylamide-bisacrylamide gels. After SDS-PAGE, proteins were transferred to nitrocellulose membranes. Membranes were blocked overnight at 4 C in 5% BSA-TTBS (0.1% Tween-20, 20 mmol/L Tris, 150 mmol/L NaCl), washed in TTBS, and incubated for 2 h with antiphosphotyrosine antibody (PY20-HRP diluted 1:1000), or with anti-KGF-R antibody (1 µg/mL ), followed by incubation with peroxidase-conjugated secondary antibody (diluted 1:5000). Finally, probed proteins were revealed by an enhanced-chemiluminescence system (BM). After the first blotting with peroxidase-conjugated antiphosphotyrosine antibody, nitrocellulose membranes were stripped at 50 C for 30 min in stripping buffer (100 mmol/L 2ß-mercaptoethanol, 2% sodium dodecyl sulphate, 62.5 mmol/L Tris-HCl, pH 6.7) and reprobed with anti-KGF antibody.
Evaluation of intracellular calcium concentration
For evaluation of intracellular calcium concentration, [Ca2+]i, cells were grown at confluence on plastic coverslips (Aclar; Allied Engineering Plastic; Pottsville, PA). During the 24 h before the experiments, cells were maintained in serum-free medium. [Ca2+]i was determined using the calcium-sensitive dye Fura-2/AM, as described previously (18). Briefly, BPH cells were loaded with 2 µmol/L Fura-2/AM for 45 min at 37 C, washed, resuspended in serum-free medium, and incubated for another 20 min in Fura-2-free medium. Cells were then washed with Krebs-Heseleit HEPES-KHH buffer (pH 7.4), containing 1.25 mmol/L CaCl2, 5.36 mmol/L KCl, 0.81 MgSO4, 130.62 mmol/L NaCl, 5.55 mmol/L glucose, 8.60 mmol/L HEPES sodium salt, and 11.7 mmol/L HEPES free acid. Coverslips were then mounted diagonally in a quartz cuvette so that the excitation and emission paths were at 45° angle to the coverslip. The cuvette, containing 2 mL HKK buffer, was maintained at 37 C. Fluorescence was measured using a spectrofluorometer (University of Pennsylvania Biomedica Group, Philadelphia, PA) with a single-wavelength excitation (340 nm)/emission (510 nm). Stimuli were added directly in the cuvette. Calibration was performed using ionomycin (0.02 mmol/L) to obtain maximum fluorescence, followed by EGTA (8 mmol/L) to obtain minimum fluorescence. Fluorescence measurements were converted to [Ca2+]i, according to Grynkiewicz et al. (19), assuming a dissociation constant of Fura-2 for calcium of 224 nmol/L.
Electron microscopy
Transmission electron microscopy (TEM). Cells were seeded onto tissue culture dishes (10-cm diameter) and stimulated for 48 h with analogue (V) alone or in presence of KGF, at the same concentrations as described above. TEM was performed on cell cultures, as previously described (20). Briefly, after trypsinization, the cell pellets, obtained by centrifugation at 800 x g for 5 min, were fixed in 2.5% glutaraldehyde, 0.1% paraformaldehyde in 0.1 mol/L cacodylate buffer (pH 7.4) at room temperature for 1 h, and postfixed with 1% osmium tetroxide in the same buffer for 1 h. After dehydration, cell pellets were embedded in Epon 812 (Fluka Chemical Co., Buchs, Switzerland). Ultra-thin sections were stained with uranylacetate, followed by lead citrate, and examined with a Phillips 410 electron microscope.
Scanning electron microscopy (SEM). SEM was performed on cell cultures, as previously described (20). Cells were grown on sterile glass coverslips and treated as described for TEM. Subsequently, cultures were fixed with 1.5% glutaraldehyde in 0.1% cacodylate buffer at room temperature for 24–36 h. After rinsing in the same buffer, samples were dehydrated with progressive acetone dilutions and subjected to critical point drying with CO2. Cells were then coated with gold-palladium in a 5001 cool Polaron sputtering apparatus, mounted on stubs, and observed with an S 400 F.E. scanning electron microscope (Hitachi)operated at 15–20 Kv.
In situ end labeling (ISEL)
BPH cells (104 cells/mL ) were seeded on sterile glass slides in their growth medium into 150-mm diameter culture dishes. Cells were accurately washed in PBS and incubated in the same conditions as previously described. Apop Tag in situ apoptosis detection kit peroxidase was used to detect the presence of DNA strand breaks in apoptotic cells, following the manufacturer’s instruction. It performs a nonisotopic DNA end extension in situ and immunohistochemical staining of the extended DNA. Residues of digoxigenin-11–2',3'-dideoxy-uridine-5'triphosphate are catalytically added to the 3'-OH ends of double- or single-stranded DNA by terminal transferase (TdT). Antidigoxigenin antibodies, carrying a conjugated reporter enzyme (peroxidase) to the reaction site, detect the incorporated nucleotides. The localized peroxidase enzyme then catalytically generates an intense signal from chromogenic substrate (diaminobenzidine). Incubating cells in the absence of TdT enzyme performed the control for method specificity. The percentage of apoptotic cells (the number of stained cells divided by the total number of cells) was calculated in at least five separate fields per slide in five different slides.
A trypan blue exclusion assay was performed, to rule out toxic effects of analogue (V) on BPH cells (n = 2 in two distinct preparations of BPH cells).
Immunocytochemical determination of bcl-2 expression
The immunocytochemical staining procedure was performed as previously described (21). Briefly, the cells were seeded on sterile glass slides in their growth medium, into 150-mm-diameter culture dishes. Near to confluence, cells were accurately washed in PBS and incubated in phenol red- and serum-free medium containing 0.1% BSA with analogue (V) (10 nmol/L) and/or KGF (10 ng/mL ), for 48 h. Slides were washed twice with PBS (pH 7.4) and fixed in 3.7% paraformaldehyde in PBS for 15 min at room temperature, followed by permeabilization in 3.7% paraformaldehyde in PBS, containing 0.1% Triton X-100, for 15 min at room temperature. Antihuman bcl-2 mouse monoclonal antibody was diluted in PBS (1:40) containing 2% BSA, added to the slides, and incubated overnight at 4 C. Slides were washed three times (5 min) in PBS and incubated at room temperature for 45 min with 2% BSA-PBS, containing the second antibody (dilution 1:1000). After washing three times in PBS, the slides were examined with a phase-contrast microscope (mocrophot-FX microscope; Nikon, Kogaku, Tokyo, Japan). Slides lacking the primary antibody or stained with the corresponding nonimmune serum were processed as controls.
The percentage of bcl-2 expression (the number of stained cells divided by the total number of cells) was calculated in at least five separate fields per slide in five different slides.
Statistical analysis
Statistical analysis was performed by one-way ANOVA and unpaired Student’s t tests, when appropriate. The computer program ALLFIT (22) was used for the analysis of sigmoidal dose-response curves to obtain estimates of half-maximal inhibition values (IC50). Data were expressed as mean (±SE).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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As shown in Fig. 1
, incubation of
BPH cells with KGF (0.1–100 ng/mL) resulted in a significant increase
in the proliferation rate at all the concentrations tested
(P < 0.01 vs. control; n = 2, in two
distinct preparations of BPH cells). Maximal effect was obtained at the
concentration of 10 ng/mL (1.8-fold increase). The stimulatory effect
of increasing concentrations (1–100 ng/mL ) of KGF was completely
blunted by the simultaneous incubation with a fixed concentration (10
nmol/L) of vitamin D3 (P < 0.01
vs. KGF-treated cells; n = 2, in two distinct
preparations of BPH cells, Fig. 1). Similar results were obtained with
an equimolar concentration of the vitamin D3
analogue, analogue (V), (P < 0.01 vs.
KGF-treated cells; n = 3, in two distinct preparations of BPH
cells, Fig. 1). As shown in Fig. 1, incubation of BPH cells with 10
nmol/L of either vitamin D3 or its analogue
induced a significant inhibition in cell growth, when compared with
their relative controls (P < 0.01 vs.
control). Even in KGF-treated cells, coincubation with vitamin
D3 or analogue (V) significantly reduced cell
proliferation below the basal control (P < 0.01
vs. untreated cells). Therefore, we tested the effect of
increasing concentrations of vitamin D3 and
analogue (V) on unstimulated BPH cells. Results are reported in Fig. 2. Both vitamin D3
(Fig. 2A, n = 2, in two separate preparations of BPH cells) and
analogue (V) (Fig. 2B, n = 3, in three separate preparations of
BPH cells) dose-dependently inhibited cell growth, with
IC50s = 5.4 ± 2.4 and 2.5 ± 1.6
nmol/L, respectively. The effect was significant, even in subnanomolar
concentrations (P < 0.01 vs. control).
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Because the incubation with vitamin D3 or
its analogue induced a consistent delay in cell proliferation, we
investigated whether or not these compounds induce cell death.
Preliminary experiments performed using the trypan blue exclusion assay
indicated that the percentage of viable cells is not different in
cultures treated, or not, with the analogue (V) (10 nmol/L), up to
48 h (not shown). Hence, analogue (V) is not apparently toxic for
BPH cells. However, TEM analysis of BPH cells indicated that treatment
with analogue (V) (10 nmol/L, 48 h) induced the typical
ultrastructural features of programmed cell death, apoptosis (Fig. 3A). The morphological features of
apoptosis included sharply compacted masses of chromatin (as a result
of nuclear fragmentation), cytoplasmatic condensation, and closely
packed (but well-preserved) mitochondria. A micrograph of control BPH
cells is in Fig. 3B. Figure 3D shows the morphological appearance of
BPH cells, when analyzed by SEM. Treatment, as before, with analogue
(V), induced an irregular, so-called boiling shape of the cell surface,
also indicative that a death program is activated (Fig. 3C). Similar
results were also obtained with a simultaneous treatment with analogue
(V) (10 nmol/L) and KGF (10 ng/mL , 48 h), although to a lower
extent (not shown). To quantify this phenomenon, we used ISEL. Results
are reported in Table 1. After 48 h,
the fraction of highly stained apoptotic nuclei in analogue (V)-treated
cells was 50 ± 1.03%; whereas in untreated cells, it was 10
± 0.44% (P < 0.01). The simultaneous treatment of
BPH cells with analogue (V) (10 nmol/L) and KGF (10 ng/mL) relatively
decreased the number apoptotic cells (40 ± 0.62%), when compared
with analogue (V) alone (P < 0.01); however, the
number of labeled nuclei was still higher than in untreated cells
(P < 0.01). KGF (10 ng/mL ) alone reduced the number
of ISEL-positive cells (4 ± 0.28%), below the basal control
value (P < 0.01). Since bcl-2 is deeply involved in
the prevention of the activation of cell death program, we investigated
the expression of this oncoprotein after different treatments with
analogue (V) (10 nmol/L), KGF (10 ng/mL ), and their combination for
48 h (Table 1). We found that KGF stimulates bcl-2 expression over
the basal control value (P < 0.01), whereas the
simultaneous addition of analogue (V) partially prevents this effect
(P < 0.01). Treatment with analogue (V) decreased the
number of bcl-2-positive cells below the control value
(P > 0.01).
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When BPH cell cultures were incubated for 5 min, in the presence
of KGF (10 ng/mL ), there was an evident increase in tyrosine
phosphorylation of a 120-kDa protein (Fig. 4A). Simultaneous treatment of BPH cells
with analogue (V) (10 nmol/L) and KGF (10 ng/mL ) reduced the increase
in tyrosine phosphorylation of this protein, whereas analogue (V) alone
was without effect (Fig. 4A). The same results were obtained after 10
and 15 min of incubation (data not shown). After stripping, incubation
of the same membrane with an anti-KGF-R antibody revealed that the
120-kDa protein band corresponded to the KGF-R (Fig. 4B). Figure 5 shows the typical effect of analogue
(V) on [Ca2+]i in Fura
2-loaded BPH cells. In three separate experiments, analogue (V) induced
a rapid and sustained increase in
[Ca2+]i (2.1 ±
0.3-fold increase over the basal level).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In this study, we confirmed that KGF is a potent mitogen in BPH cells (15) and originally demonstrated that KGF prevents naturally occurring prostatic cell apoptosis by stimulating bcl-2 protein expression. This protein is a member of the anti- and proapoptotic factor family that regulates the susceptibility to programmed cell death. In particular, bcl-2 availability blocks the process of cell death (27) and, therefore, it is considered a survival factor also involved in development and progression of hormone refractory prostate cancer (28, 29). Hence, KGF might induce prostate hyperplasia by a dual mechanism of action: stimulating cell proliferation, and inhibiting cell death.
This study demonstrates, for the first time, that vitamin D3, or one of its analogues, partially counteracts KGF-mediated effects in BPH cells. Indeed, vitamin D3 and/or analogue (V) not only blunted KGF-induced cell proliferation but also reduced KGF-stimulated expression of bcl-2 protein, thereby partially restoring programmed cell death in BPH cells. It is important to note that interaction among vitamin D3, or its analogue, and KGF seems to occur at the level of the receptor tyrosine phosphorylation. In fact, analogue (V) completely prevents KGF-induced tyrosine phosphorylation of a 120-kDa protein, corresponding to the KGF-R.
In this study, we also found that nanomolar concentrations of vitamin D3 and analogue (V) decreased cell proliferation. Although this effect might be related to a decreased progression of BPH cells into the cell cycle, it might also be mediated by induction of apoptosis and by a decreased expression of the bcl-2 protein. The proapoptotic activity of analogue (V) could be attributable to the induced increase in intracellular calcium concentration, because in several systems, an overloaded intracellular calcium is linked to cell death (30, 31, 32).
Finding that the analogue (V) transiently alters intracellular calcium dynamics is not surprising, because similar effects of vitamin D3 have been described in other cell types as chondrocyte (33), osteoblast (34), and parathyroid cells (35). This rapid effect on intracellular calcium suggests that analogue (V) transduces, at least in part, its action in BPH cells via a nongenomic mechanism. Plasma membrane receptors for vitamin D3 or its metabolite has been suggested by several Authors (35, 36, 37, 38, 39) and recently demonstrated by Nemere et al., 1998 (40), and Pedrozo et al., 1999 (41), in chondrocytes. These membrane receptors bind vitamin D3 in subnanomolar concentrations and are not only coupled to intracellular calcium mobilization but also to an increase in inositol trisphosphate and diacylglycerol levels (34), with activation of protein kinase C (42, 43) via a phospholipase C-dependent mechanism (44). Because genistein, a tyrosine kinase inhibitor, did not block the vitamin D3-induced activation of protein kinase C in chondrocytes, it has been suggested that tyrosine phosphorylation is not involved in this pathway (41). Our results with analogue (V) in BPH cells further support the concept that vitamin D3 does not induce tyrosine phosphorylation, and they indicate a novel mechanism of action for these molecule(s) via inhibition of KGF-induced phosphorylation of a 120-kDa protein, corresponding to the KGF-R. Whether or not this effect is mediated by a nongenomic membrane receptor needs to be established; however, the rapid effect itself suggests that this might be the case.
In conclusion, according to a previous study (15), we report that KGF is a potent mitogen for BPH cells, and we speculate that this growth factor might be involved in the pathogenesis of BPH. Because analogue (V) completely counteracts KGF effects in human BPH cells, we suggest it as a new candidate for the therapy of this common male disorder.
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Acknowledgments |
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Footnotes |
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Received August 10, 1999.
Revised April 9, 2000.
Accepted April 13, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-reductase inhibitor on prostate tissue androgens and
prostate-specific antigen. J Clin Endocrinol Metab. 71:1552–1555.[Abstract]
-reductase inhibitor, finasteride, on benign prostatic hyperplasia.
The Finasteride Study Group. J Urol. 147:1298–1302.[Medline]
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