Activation of Caspases-3, -6, and -9 during Finasteride Treatment of Benign Prostatic Hyperplasia
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《临床内分泌与代谢杂志》
Abstract
Benign prostatic hyperplasia (BPH) results from an increase in both epithelial and stromal compartments of the human prostate. Although inhibitors of 5-reductase such as finasteride have been shown to reduce the size of BPH tissues by inducing apoptosis, their mechanisms of action still remain unknown. The present study supports that such a process triggered by finasteride is caspase dependent with a possible involvement of two effector caspases (caspase-3 and 6) and two initiator caspases (caspase-8 and 9). Indeed, by using tissues from patients affected by BPH and treated by finasteride (5 mg/d) for 2–3, 6–8, or 27–32 d, we observed that the 5-reductase inhibitor induced apoptosis in epithelial cells (evaluated through cell number positive for terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end labeling) as early as 2–3 d of treatment, with a maximal activity (250-fold increase, P < 0.0001) at 6–8 d of treatment. However, after 27–32 d of treatment, the number of apoptotic cells was reduced and was close to control. Caspases-3, -6, -8, and -9 were immunolocalized to (basal and secretory) epithelial cells and to a lesser extent to stromal cells. Activated caspase-3 immunoexpression was restricted to epithelial secretory cells, and its immunostaining intensity appeared to be higher in BPH tissues from patients treated for 2–3 or 6–8 d. Consistently, in Western blotting analyses, activated caspases-3 and -6 were detected as early as 2–3 d of treatment in BPH tissues, and their levels were increased after 6–8 d of treatment. In real time quantitative PCR experiments, caspase-3 and -6 mRNA levels were found to be unchanged after finasteride treatment. Activated caspase-8 was not detected in the different conditions tested, whereas activated caspase-9 protein levels were maximally enhanced after 2–3 d of finasteride treatment. In conclusion, we report here that finasteride treatment of BPH tissues induced a caspase-dependent apoptotic process restricted to epithelial cells by activating effector caspases-3 and -6 and exhibited a transient action because the apoptotic process was no longer observed after 27–32 d of treatment.
Introduction
BENIGN PROSTATIC HYPERPLASIA (BPH) is an extremely common pathology among aging men. Indeed, approximately 85% of men over age 50 yr will develop BPH. By their 90s, half of these men will require treatment for symptomatic relief of urinary obstructive symptoms associated with BPH. Urinary obstructive symptoms have both an anatomic component of obstruction produced by the enlarging hyperplastic tissue and a dynamic component related to smooth muscle tone (1). Histological studies have shown that hyperplasia results from an increase of the epithelial and stromal components of the prostate (2, 3).
The progression of prostate gland to BPH may rely on androgen action (4). Androgens are key hormones during prostate development. Indeed, androgen deprivation results in either a maldevelopment of the prostate or a reduction in prostate size. The androgen mainly responsible for these events is dihydrotestosterone (DHT), which results from the conversion of testosterone by the 5-reductase enzyme in prostate stromal and epithelial cells. DHT is primarily responsible for prostate development and is also supposed to play a major role in the pathogenesis of BPH (5, 6), suggesting that inhibition of DHT formation may exert a beneficial effect on BPH development. Two types of 5-reductase enzymes have been identified. 5-Reductase type I is predominant in extraprostatic tissues, such as skin and liver. Type II is the predominant enzyme expressed in prostatic stromal and epithelial cells (4). Different drugs, such as finasteride, have been generated to inhibit the 5-reductase activity, particularly the type II. Finasteride administration induces a 85–90% decrease in DHT levels within the human prostate concomitantly with a 20–30% reduction of prostatic size (7, 8, 9, 10). Although histological changes related to an apoptotic process within the prostates of patients treated with finasteride have been reported (11), the molecular mechanisms underlying such a process still remain to be investigated.
Apoptosis is a regulated process of autodigestion of cells, which involves the disruption of cytoskeletal integrity, cell shrinkage, nuclear condensation, and the activation of endonucleases. The chief effectors of the apoptotic cell death pathway are the caspase family of cysteine proteases. Caspases are synthesized as inactive precursors (procaspases) that must be cleaved at specific aspartate residues to generate the active (cleaved) subunits. Procaspase cleavage can occur by several mechanisms including proximity-induced autoprocessing or cleavage by other caspases, revealing a caspase cascade with upstream initiator caspases such as caspases-8, -9, and -10 and downstream effector caspases such as caspases -3, -6, and -7. The activity of these caspases is regulated by several families of both pro- and antiapoptotic cellular proteins (for reviews, see Refs.12 , 13).
In this study, in BPH tissues from patients treated for different periods with finasteride, we addressed the question as to whether the apoptotic process is caspase dependent. For this purpose, we immunolocalized and examined at different periods of finasteride treatment the expression and activation of two effector caspases (caspase-3, -6) and two initiator caspases (caspase-8, -9).
Subjects and Methods
A retrospective analysis was performed on patients treated or not with finasteride at the Department of Urology at Centre Hospitalier Lyon-Sud. According to the length of finasteride (5 mg/d) treatment, four groups of patients were determined: untreated (control) patients (n = 10), patients treated for 2–3 d (n = 7), patients treated for 6–8 d (n = 8), and patients treated for 27–32 d (n = 4). A total of 29 patients were included in this study. The mean age of the patients was 72 yr (range 58–93 yr). Characteristics of the patients in terms of age, plasma prostate-specific antigen (PSA) and prostate volume before finasteride treatment are presented in Table 1. The different patient groups are homogeneous because no statistical differences for the different parameters (age, PSA, prostate volume) among the groups were observed. Patients taking other medication known to possibly influence androgen levels were excluded. Endourethral resection was performed 2–3, 6–8,or 27–32 d after treatment. The fragments were either fixed in Bouin’s liquid or frozen in liquid nitrogen. All the cases were reviewed by a pathologist (M.Dec.), and the diagnosis of BPH was confirmed. For each case, a representative block of the BPH tissues was chosen for the immunohistochemical study. The ethics committee of the medical faculty and the state medical board agreed to these investigations, and informed consent was obtained for all patients.
Materials
Horseradish peroxidase-labeled antirabbit IgG and chemiluminescent kit were obtained from CovalAb (Lyon, France). Horseradish peroxidase-labeled antigoat IgG (SC-2020), rabbit polyclonal antibody raised against caspase-9, which reacts with both procaspase-9 (Mr 35,000) and cleaved caspase-9 (Mr 24,000), goat polyclonal antibody raised against caspase-8, which reacts with both procaspase-8 (Mr 55,000) and cleaved caspase-8 (Mr 25,000), and rabbit polyclonal antibody raised against caspase-3, which reacts with both procaspase-3 (Mr 30,000) and cleaved caspase-3 (Mr 17,000) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal antibody raised against cleaved caspase-3, which detects only the large fragment of activated caspase-3 (Mr 17,000), and rabbit polyclonal antibody raised against caspase-6, which detects procaspase-6 (Mr 34,000) and activated caspase-6 (Mr 12,000), were obtained from Ozyme (Saint Quentin en Yvelines, France). Mouse monoclonal keratin-903 antibody was obtained from Enzo Diagnostics, Inc. (Farmingdale, NY). The Envision+ kit (mouse or rabbit), the LSAB kit, Meyer’s hematoxylin, and Faramount were obtained from Dako (Trappes, France). Superfrost plus glass slides were obtained from Menzel-Glaser (Frelburg, Germany). Thymus terminal deoxynucleotidyl transferase, biotin deoxyuridine 5-triphosphate (dUTP), LC FastStart master SYBR green kit, and LC capillaries were obtained from Roche Molecular Biochemicals (Mannheim, Germany). Cobalt chloride, sodium cacodylate, extravidin peroxidase, diaminobenzidine (DAB), nickel chloride, and random primers were purchased from Sigma (Meylan, France). Finasteride (Proscar) was obtained from MSD (Whitehouse Station, NJ). Dithiothreitol, deoxynucleotide triphosphates, and Moloney murine leukemia virus reverse transcriptase were obtained from Invitrogen SARL (Cergy Pontoise, France).
Methods
Terminal deoxynucleotidyl transferase-mediated dUTP nick labeling (TUNEL). Paraffin sections (5 μm) of Bouin-fixed prostatic tissues were mounted onto Superfrost plus slides. The sections were deparaffinized and rehydrated (xylene 5 min, ethanol 100, 95, 70%, 30 sec each) and then washed in distilled water before beginning the TUNEL reaction. The slides were transferred to a plastic jar containing 0.01 M citrate buffer (pH 6) and microwave irradiated for 5 min (370 W) and then left 20 min at room temperature (RT). After a wash with phosphate buffer saline (PBSx 1), endogenous peroxidases were blocked for 5 min with H2O2 2%. Sections were washed three times with PBSx 1. The specimens were then incubated 60 min at 37 C in a moist chamber with TUNEL mix, which consisted of 0.3 U/μl calf thymus terminal deoxynucleotidyl transferase, 0.007 nmol/μl biotin dUTP, 1 mM cobalt chloride, 30 mM Tris (pH 7.2), and 140 mM sodium cacodylate. After washing (four PBS baths of 5 min each at RT) the sections were saturated 10 min with BSA 2% at RT. Sections were treated for 30 min at 37 C in moist chamber with a 1:20 dilution of Extravidin peroxidase. After three washes in PBS, detection was performed with DAB (1.25 mg), 25 μl nickel chloride 3%, 152 μl Tris-HCl 1 M (pH 7.5), completed with distilled water to 2 ml). Slides were mounted in Faramount.
SYBR green real-time PCR. Total RNAs were extracted from BPH tissues with TRIzol reagent (untreated patients, n = 7; patients treated with finasteride for 2–3 d, n = 4; for 6–8 d, n = 4; for 28–32 d, n = 6). cDNAs were obtained from 1 μg RNA incubated 1 h at 37 C in the presence of 50 μM random primers, 200 μM deoxynucleotide triphosphates, 100 mM dithiothreitol, and 200 U/μl of Moloney murine leukemia virus in a total volume of 20 μl.
In a first step, we assessed standards for caspase (-3 and -6) and ?-actin RNA quantitation by SYBR green PCR. The standards were assessed by scalar dilution of caspase or ?-actin cDNA. The samples were run in triplicate.
SYBR green real-time PCR assays were carried out in 20 μl PCR mixture volume consisting of 2 μl of LC FastStart SYBR green master mix, containing HotStarTaq DNA polymerase, 1 μl of 10 μM of each oligonucleotide primer, 4 μl of diluted RT mix (1/40) and 5 mM MgCl2.
Caspase-3 or -6 or ?-actin gene amplification was carried out as follows: initial activation of HotStarTaq DNA polymerase at 95 C for 480 sec; 45 cycles in three steps: 95 C for 15 sec, 55 C (caspase-3), 59 C (caspase-6), or 68 C (?-actin) for 6 sec, and 72 C for 12 sec. At the end of amplification cycles, melting temperature analysis was carried out by a slow increase in temperature (0.1 C/sec) up to 95 C. The primers used were: caspase-3, 5'-GAGCTGCCTGTAACTTG-3' and 5'-ACCTTTAGAACATTTCCACT-3'; caspase-6, 5'-ACTGGCTTGTTCAAAGG-3' and 5'-CAGCGTGTAAACGGAG-3’; and ?-actin (14). PCR products were checked by direct sequencing. Amplification, data acquisition, and analysis were carried out by LightCycler instrument (Roche Molecular Biochemicals) using LightCycler software (version 5.3.2, Roche). This software, coupled to the LightCycler instrument, determines the threshold cycle that represents the number of cycles in which the fluorescence intensity is significantly above the background fluorescence. The amounts of caspase (-3 or -6) relative to the amount of ?-actin in the same sample were analyzed by the relative quantification approach with the use of efficiency correction using Real Quant 1.0 software (Roche Molecular Biochemicals). The results are expressed as a normalized ratio.
Western blotting analysis. Proteins were prepared from tissue samples frozen in liquid nitrogen and then stored at –70 C until use. Prostatic tissues were homogenized in 200 μl ice-cold hypotonic buffer [25 mM Tris-HCl (pH 7.4), protease inhibitor cocktail] and sonicated (10 sec at 80). Protein concentration was determined by the Bradford assay. For Western blot, three patients for each experimental condition were used. The same antibodies were used for immunohistochemistry and Western blot analyses.
Proteins (100 μg) were resolved on 10% sodium dodecyl sulfate/polyacrylamide gels and electrophoretically transferred to nitrocellulose membranes using Tris 15 mM-glycine 120 mM buffer (pH 8.3) containing 20% methanol at a constant voltage of 100 V for 30 min. Under those conditions, the transfer of low-molecular-weight proteins is favored, i.e. transfer of activated caspases (which Mr ranges from 12,000 to 20,000). After transfer, the membranes were incubated in a blocking buffer [Tris-buffered saline (TBS) containing 5% fat-free dry milk and 0.1% Tween 20] 2 h at RT, rinsed three times with TBS/0.1% Tween 20 (3 x 10 min), and then incubated with the first antibody (in TBS containing 2% fat-free dry milk) overnight at 4 C. Anti-caspase-3 and anticleaved caspase-3 were diluted at 1/100, anticaspase-6 was diluted at 1/200, and anticaspase-8 and -9 were diluted at 1/100. The membranes were then rinsed with TBS/0.1% Tween 20 (3 x 10 min) and incubated with horseradish peroxidase-labeled antirabbit IgG (1/2000) or antigoat IgG (1/3000) in TBS containing 1% fat-free dry milk and 0.1% Tween 20 for 1 h at RT. The membranes were thoroughly washed with TBS/0.1% Tween 20 (3 x 10 min) and then with TBS. Bound antibodies were detected by the chemiluminescence kit and Biomax MR films (Sigma, Meylan, France). Protein loading was checked by reprobing the blot with a rabbit anti-actin IgG (1/500). The Biomax MR films were scanned on a Gel doc 2000 apparatus (BioRad Life Science Group, Marnes La Coquette, France), and quantitation was realized with Quantity one Software (Bio-Rad Life Science Group). The results are expressed as a ratio of caspase to actin.
Immunohistochemistry. Paraffin sections of Bouin-fixed prostatic tissues were sectioned at 5 μm. The sections were mounted on Superfrost plus glass slides, deparaffinized, hydrated, treated 20 min at 93–98 C in citric buffer [0.01 M (pH 6)], rinsed in osmosed water (2 x 5 min), and washed (2 x 5 min) in TBS (pH 7.8). For anticaspase-3 (pro and cleaved), anticleaved caspase-3, anticaspase-6, and anticaspase-9 antibodies, the Envision plus kit was used. For the anticaspase-8 antibody, the LSAB kit was used. The first steps were similar for both kits. Briefly, endogenous peroxidases were blocked with 3% H2O2 for 15 min and then washed three times in TBS. The sections were then incubated overnight at 4 C with the primary antibody diluted in antibody diluent (anticaspase-3, 1/100; anticleaved caspase-3, 1/50; anticaspase-6, anticaspase-9, and anticaspase-8, 1/100). The sections were then washed (2 x 5 min) in TBS. For the Envision plus kit, the sections were incubated 30 min at 37 C in presence of the secondary antibody peroxidase-conjugated. The sections were then rinsed (2 x 5 min) in TBS, incubated 10 min at RT with 3-amino-9-ethylcarbazole or DAB, which generated a red color at the site of peroxidase activity and rinsed (2 x 5 min) in osmosed water. For the LSAB kit, after incubation with the primary antibody, the section were incubated with the biotinylated secondary antibody, and after washes, a peroxidase streptavidin complex was applied. DAB or 3-amino-9-ethylcarbazole was used as a peroxidase chromogen. For double-staining experiments, the slides were first incubated with the keratin-903 primary antibody (1/200). After washes, the slides were incubated with an antimouse secondary antibody peroxidase-conjugated (mouse Envision plus kit) and revealed with DAB. After washing, the slides were incubated with the anticleaved caspase-3 antibody (1/50). The antigen-antibody complexes were revealed with the LSAB kit with a streptavidin phosphatase conjugate and revealed with fast red. For the different experiments, sections were counterstained with Meyer’s hematoxylin. Finally, sections were mounted in Faramount. As a negative control, the primary antibody was replaced by the antibody diluent. Immunohistochemistry was performed on a slide from each patient. For analysis of immunohistochemical staining, the intensity was rated as none, weak, moderate, or intense for each slide. Specimens were considered immunopositive when 1% or more of the BPH tissue had clear evidence of immunostaining. The immunostaining was evaluated by two independent observers in the laboratory, blinded as to the treatment status. Similar results were obtained for the different patients; a representative immunohistochemistry is presented.
Data analysis. Depending on the availability of BPH tissues, the number of patients studied in the different groups is as follows: four to 10 patients per condition for the immunohistochemical experiments, three patients per condition for the Western blotting analyses, and four to seven per condition for the real-time PCR experiments. For statistical analysis one-way ANOVA was performed to determine whether there were differences among all groups (P < 0.05), and then the Bonferroni posttest was performed to determine the significance of the differences between the pair of groups. P < 0.05 was considered significant. The statistical tests were performed on StatView software (version 5.0, SAS Institute Inc., Cary, NC).
Results
Time dependency of the apoptotic process in BPH tissues on finasteride administration
Very few TUNEL-positive (apoptotic) cells were observed in BPH tissues from untreated patients (Fig. 1A). In contrast, a higher amount of TUNEL-positive cells were visualized in epithelial secretory cells of prostatic tissues from patients affected by BPH and treated during 2–3 or 6–8 d with finasteride (Fig. 1B). A very limited number of apoptotic cells was observed in stromal cells (Fig. 1, A and B).
Consistently, the evaluation of the number of TUNEL-positive cells in BPH tissues indicated an increase in apoptotic cells after 2–3 d of treatment and a more dramatic increase after 6–8 d (250-fold, P < 0.0001). However, after 27–32 d of finasteride treatment, the apoptotic cell number decreased and was back to control values (Fig. 1C).
To further characterize the apoptotic cell death process in prostatic epithelial cells, four caspases, two effector caspases (caspase-3 and -6) and two initiator caspases (caspase-9 and –8), were studied in terms of proteins in BPH tissues from untreated or finasteride-treated patients.
Immunolocalization of caspases-3, -6, -9, and -8 in BPH tissues after finasteride treatment
The immunohistochemical experiments were performed in BPH tissues from each patient, treated or not treated with finasteride. A representative experiment is presented for each different caspase. For the immunolocalization of caspase-3, two different antibodies were used. The first one detected both pro- and cleaved caspase-3 (Fig. 2, A–D), whereas the second one detected exclusively cleaved (activated) caspase-3 (Fig. 2, E–H). In tissues from untreated patients, caspase-3 was mainly detected in basal and secretory epithelial cells and to a lesser extent in stromal cells (Fig. 2A). After finasteride treatment, caspase-3 immunolocalization in BPH tissues was not affected (Fig. 2), although an apparent increase in caspase-3 immunostaining intensity, depending on the different times of treatment including 2–3 d (Fig. 2B) and 6–8 d (Fig. 2C), could be observed. Interestingly, cleaved caspase-3 immunostaining was either not detected or detected at very low levels in BPH tissues from untreated patients (Fig. 2E), whereas, in contrast, after treatment with finasteride for 2–3 and 6–8 d, its staining increased and appeared exclusively in the epithelial secretory cells (Fig. 2, F and G). Then 27–32 d after finasteride treatment, the cleaved caspase-3 staining intensity was reduced and appeared to be similar to that observed in tissues from untreated patients (Fig. 2H). To further evaluate whether activated caspase-3 was expressed in the secretory or basal epithelial cells of prostate, we used an antibody raised against keratin-903, known to be expressed exclusively in the basal epithelial cells. Keratin-903 staining was observed in the basal epithelial cells of BPH tissues from both untreated patients and patients treated with finasteride for 2–3, 6–8, and 28–32 d (Fig. 2, I–L). Activated caspase-3 staining was detected (exclusively) in secretory epithelial cells but not in basal cells of BPH tissues from patients treated with finasteride for 2–3 and 6–8 d (Fig. 2, J and K). In BPH tissues from patients treated with finasteride for 28–32 d, prostatic secretory epithelial cells remained detectable, although activated caspase-3 staining was no longer observed (Fig. 2L). Such an observation suggests that the decrease in activated caspase-3 staining in tissues from patients treated with finasteride for 28–32 d was probably not related to epithelial cell loss.
For the immunolocalization of caspase-6, the antibody used recognized both pro- and cleaved caspase-6. In BPH tissues from both control untreated and treated patients, caspase-6 was immunolocalized to secretory and basal cells and to a lesser extent to stromal cells (Fig. 3A). However, an apparent increase in the intensity of caspase-6 staining in the epithelial secretory cells could be observed in tissues from patients treated for 2–3 and 6–8 d with finasteride (Fig. 3, B and C). After 27–32 d of treatment, the caspase-6 staining intensity appeared to be reduced and was back to the staining intensity observed in tissues from untreated patients (Fig. 3D).
Antibodies used for the detection of initiator caspases (caspase-9 and -8) were designed to recognize both pro- and cleaved caspases. Caspase-8 was immunodetected in the epithelial secretory, basal, and the stromal cells from the untreated patients (Fig. 3E). The intensity of caspase-8 immunostaining appeared not to be affected by finasteride treatment in BPH tissues (Fig. 3, F–H). In BPH tissues, caspase-9 was immunoexpressed in the secretory epithelial and stromal cells (Fig. 3I). In contrast to caspase-8, the intensity of caspase-9 immunostaining appeared to be higher in BPH tissues from patients after 2–3 or 6–8 d of treatment (Fig. 3, J and K), whereas it was reduced and was similar to that observed in tissues from patients treated with finasteride for 27–32 d (Fig. 3L).
To strengthen the data obtained from the immunohistochemical experiments, we further carried out quantitative studies in BPH tissues by Western blotting analyses on caspase protein levels and real-time quantitative PCR on caspase mRNA levels.
Effects of finasteride treatment on caspase-3, -6, -8, and -9 protein levels in BPH tissues
The same antibodies used for the previous immunohistochemical studies were also used for Western blotting experiments. The Western blotting experimental conditions were optimized to detect the active-cleaved caspases (see Materials and Methods). Activated caspases [caspase-3 (Mr 17,000) and -6 (Mr 12,000)] protein levels were low in untreated patients, whereas their levels were significantly (P < 0.0003) increased after a 6–8 d of finasteride treatment (Fig. 4, A and B). Caspase-8 was not detected in terms of cleaved caspase in BPH tissues from patients treated or not with finasteride (data not shown). The cleaved caspase-9 protein (Mr 20,000) was detected at low levels in untreated patient tissues, but such levels were significantly (P < 0.0113) increased in tissues from patients treated during 2–3 and to a lesser extent during 6–8 d of treatment with finasteride (Fig. 4C).
Effects of finasteride treatment on caspase-3 and -6 mRNA levels in BPH tissues
Caspase-3 and -6 mRNA levels in BPH tissues from untreated patients or patients treated for 2–3, 6–8, or 27–32 d with finasteride were determined by using real-time quantitative RT-PCR assays. No significant changes in caspase-3 or -6 mRNA levels were observed after finasteride treatment (Fig. 5, A and B).
Discussion
Among the therapies used against BPH (at least with a prostate weight >40 g), the treatment with a 5-reductase inhibitor such as finasteride has been reported to be effective in improving both clinical symptoms and urinary flow rates as well as in reducing prostate size. Although it has been shown that BPH treated with finasteride exhibits a progressive decrease in prostatic cell size and function during the first several months of treatment (11), the molecular mechanisms underlying finasteride action remain poorly understood.
In the present study, finasteride was shown to trigger apoptosis in BPH tissues in the first week of treatment. Indeed, apoptotic epithelial prostatic cells were identified 2–3 d after finasteride treatment with the highest level of apoptotic cell number evidenced after 6–8 d of treatment. In contrast, after 27–32 d of treatment with the inhibitor, the number of apoptotic epithelial prostatic cells was back to levels comparable with those observed in tissues from untreated patients. Such observations are consistent with those reported by Rittmaster et al. (11), although these authors have used a long-term protocol (6–18 and 23–73 d and 3–48 wk of treatment), compared with our own protocol. Indeed, the peak of apoptosis after finasteride treatment was observed after 6–8 d (our study) or 6–18 d (11). Moreover, the apoptotic process was detected in the prostatic epithelial cells but not in stromal cells of BPH tissues treated with finasteride (the present study and Ref.11). Our present data suggest that the effects of finasteride treatment in the cell death process, at least evaluated through the TUNEL approach, appear to be transient. Indeed, the death process was observed after 2–3 and 6–8 d but not after 27–32 d of treatment. Second, these data confirm that the death process affects mainly epithelial cells but not stromal cells. This observation could explain that, although finasteride is effective in reducing prostate volume, such a decrease was only of 20–30% of the initial volume because it may primarily concern the epithelial cell component (4, 10, 15, 16).
To further characterize the molecular mechanisms underlying the finasteride-induced apoptotic process, we studied the caspase proteins that are key factors in the cell death machinery (for a review, see Ref.13). In this context, we characterized the expression of pro- and especially cleaved caspases in the apoptosis induced by finasteride in BPH tissues. The effector caspase-3 and -6 were shown to be expressed in secretory and basal epithelial cells and to a lesser extent in the stromal cells. Indeed, caspase-3 was immunodetected in basal and secretory epithelial cells by an antibody recognizing both pro- and activated caspase-3 (17). One of the original contributions of the present report is the demonstration that finasteride treatment induces the activation of caspases-3 and -6 during the apoptotic cell death process in BPH tissues. Indeed, an apparent increase in the immunostaining of the caspases-3 and -6 was observed when patients were treated during 2–3 or 6–8 d with the inhibitor. More specifically, activated caspase-3, which was shown to be immunolocalized exclusively in secretory but not in basal epithelial cells of BPH tissues, exhibited an increased immunostaining 2–3 and 6–8 d after treatment with finasteride. In contrast, activated caspase-3 was not detected (or very weakly) in untreated tissues and after 27–32 d of treatment with finasteride, specifically, activated caspase-3 was no longer detected (or very weakly) in the secretory epithelial cells of the prostate. The dramatic decrease of the activated caspase-3 staining was not related to a loss of epithelial secretory cells because these cells remained detectable. Furthermore, these results were confirmed by Western blotting experiments showing an increase in activated caspase-3 and -6 protein levels in BPH by 2–3 d and at higher levels by 6–8 d in treated patients. The changes in caspase-3 and -6 levels were related to alterations of protein but not of mRNA levels. Interestingly, the activation of caspase-3 and -6 was kinetically correlated with the apoptotic process in epithelial prostatic cells because the TUNEL approach shows a peak of cell death by 6–8 d in BPH patients treated with finasteride. The absence of activated caspases in stromal prostatic cells suggests that these cells probably do not use the caspase-3 pathway to trigger cell death. Some authors suggested that stromal cells of BPH may use the caspase-7 pathway to trigger apoptosis (18).
At least two pathways, the death receptor and the mitochondrial pathways, are known to activate the effector caspases via the initiator caspases. The death receptor pathway triggers the activation of the initiator caspase-8 or -10, whereas the mitochondrial pathway triggers the activation of initiator caspase-9 (19). We examined here the expression of initiator caspases belonging to each pathway, i.e. caspase-8 and -9. Caspase-8, which was detected in human prostate cancer cell lines LNCaP (20), exhibited a low immunostaining intensity in both stromal and epithelial cells in our experiments. Moreover, finasteride treatment had no effect on the caspase-8 protein expression, suggesting that finasteride-induced apoptosis in prostatic epithelial cells probably did not result from the activation of the caspase-8 pathway. However, it cannot be excluded that caspase-8 is at play in prostate cancer cell line apoptosis. Indeed, in LNCaP cell lines, TNF, and ionizing radiations induce pro- and activated caspase-8 expressions, and an addition of androgens inhibits this caspase-8 activation (21). Based on our findings, it is quite possible that by contrast to effector caspase-3 and-6, caspase-8 is not targeted by androgen action in BPH tissues. This is consistent with our recent data showing that, whereas rat castration induces, in ventral prostate, effector caspase-3 and -6 activation, it fails to affect caspase-8 expression or activation (22).
With regard to the mitochondrial pathway, we identified caspase-9 in epithelial and stromal cells in untreated BPH tissues. The immunostaining intensity of caspase-9 appeared to increase by 2–3 d and to a lesser extent 6–8 d after finasteride treatment. This observation was confirmed by Western blotting experiments, showing a maximal increase in cleaved caspase-9 protein levels after 2–3 d of treatment, suggesting that executor caspases-3 and -6 may be activated in the mitochondrial pathway. Indeed, maximal levels of activated caspase-9 were observed by 2–3 d before the peak of activated caspases-3 and -6 (observed by 6–8 d of finasteride treatment). It is of interest to mention that the possible involvement of the mitochondrial pathway during the apoptotic process in prostatic cells was also previously reported in prostate cancer cell lines. Indeed, procaspase-9 expression was detected in LNCaP cell lines (23), and withdrawal of androgens (castration) was shown to activate caspase-9 in rat prostate epithelial cells (24).
Together, finasteride triggers apoptosis exclusively in epithelial secretory cells, and its action on the apoptotic process was transient because it was limited to a short period (<27–32 d). No TUNEL-positive cells were observed after 27–32 (our present study) or 23–73 d (11) of treatment with finasteride, and effector caspase-3 was no more activated after 27–32 d of treatment with finasteride, although epithelial cells were still present because they were detectable (our present study). The molecular mechanisms underlying this arrest in the cell death process after 1 month of treatment with finasteride remain unknown. However, despite an arrest of the finasteride-dependent apoptotic process after 1 month, the inhibitor has been shown to sustain a reduced prostate volume after 1–6 yr of therapy (10, 16, 25, 26). Whether mechanisms/processes other than apoptosis could underline the BPH tissue atrophy in the long-term treatment remains to be answered.
In summary, by using BPH tissues from patients treated with finasteride during, 2–3, 6–8, or 27–32 d, we show that the antiandrogen induces apoptosis in prostatic epithelial cells within 2–3 to 6–8 d of treatment but not later (27–32 d). Such an apoptotic process is caspase dependent because it appears to result from activation of effector caspases-3 and -6, probably involving the mitochondrial pathway as evidenced by the activation of caspase 9.
Acknowledgments
We are grateful to Dr. A. Florin-McLeer for critical reading of the manuscript.
Footnotes
This work was supported by Institut National de la Santé et de la Recherche Médicale; Ministère de l’Enseignement Supérieur et de la Recherche; and la Ligue Contre le Cancer de l’Ardèche, France.
First Published Online October 26, 2004
1 A.B. and A.R. contributed equally to the work in this manuscript.
Abbreviations: BPH, Benign prostatic hyperplasia; DAB, diaminobenzidine; DHT, dihydrotestosterone; dUTP, deoxyuridine 5-triphosphate; PSA, prostate-specific antigen; RT, room temperature; TBS, Tris-buffered saline; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick labeling.
Received April 22, 2004.
Accepted October 19, 2004.
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Benign prostatic hyperplasia (BPH) results from an increase in both epithelial and stromal compartments of the human prostate. Although inhibitors of 5-reductase such as finasteride have been shown to reduce the size of BPH tissues by inducing apoptosis, their mechanisms of action still remain unknown. The present study supports that such a process triggered by finasteride is caspase dependent with a possible involvement of two effector caspases (caspase-3 and 6) and two initiator caspases (caspase-8 and 9). Indeed, by using tissues from patients affected by BPH and treated by finasteride (5 mg/d) for 2–3, 6–8, or 27–32 d, we observed that the 5-reductase inhibitor induced apoptosis in epithelial cells (evaluated through cell number positive for terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end labeling) as early as 2–3 d of treatment, with a maximal activity (250-fold increase, P < 0.0001) at 6–8 d of treatment. However, after 27–32 d of treatment, the number of apoptotic cells was reduced and was close to control. Caspases-3, -6, -8, and -9 were immunolocalized to (basal and secretory) epithelial cells and to a lesser extent to stromal cells. Activated caspase-3 immunoexpression was restricted to epithelial secretory cells, and its immunostaining intensity appeared to be higher in BPH tissues from patients treated for 2–3 or 6–8 d. Consistently, in Western blotting analyses, activated caspases-3 and -6 were detected as early as 2–3 d of treatment in BPH tissues, and their levels were increased after 6–8 d of treatment. In real time quantitative PCR experiments, caspase-3 and -6 mRNA levels were found to be unchanged after finasteride treatment. Activated caspase-8 was not detected in the different conditions tested, whereas activated caspase-9 protein levels were maximally enhanced after 2–3 d of finasteride treatment. In conclusion, we report here that finasteride treatment of BPH tissues induced a caspase-dependent apoptotic process restricted to epithelial cells by activating effector caspases-3 and -6 and exhibited a transient action because the apoptotic process was no longer observed after 27–32 d of treatment.
Introduction
BENIGN PROSTATIC HYPERPLASIA (BPH) is an extremely common pathology among aging men. Indeed, approximately 85% of men over age 50 yr will develop BPH. By their 90s, half of these men will require treatment for symptomatic relief of urinary obstructive symptoms associated with BPH. Urinary obstructive symptoms have both an anatomic component of obstruction produced by the enlarging hyperplastic tissue and a dynamic component related to smooth muscle tone (1). Histological studies have shown that hyperplasia results from an increase of the epithelial and stromal components of the prostate (2, 3).
The progression of prostate gland to BPH may rely on androgen action (4). Androgens are key hormones during prostate development. Indeed, androgen deprivation results in either a maldevelopment of the prostate or a reduction in prostate size. The androgen mainly responsible for these events is dihydrotestosterone (DHT), which results from the conversion of testosterone by the 5-reductase enzyme in prostate stromal and epithelial cells. DHT is primarily responsible for prostate development and is also supposed to play a major role in the pathogenesis of BPH (5, 6), suggesting that inhibition of DHT formation may exert a beneficial effect on BPH development. Two types of 5-reductase enzymes have been identified. 5-Reductase type I is predominant in extraprostatic tissues, such as skin and liver. Type II is the predominant enzyme expressed in prostatic stromal and epithelial cells (4). Different drugs, such as finasteride, have been generated to inhibit the 5-reductase activity, particularly the type II. Finasteride administration induces a 85–90% decrease in DHT levels within the human prostate concomitantly with a 20–30% reduction of prostatic size (7, 8, 9, 10). Although histological changes related to an apoptotic process within the prostates of patients treated with finasteride have been reported (11), the molecular mechanisms underlying such a process still remain to be investigated.
Apoptosis is a regulated process of autodigestion of cells, which involves the disruption of cytoskeletal integrity, cell shrinkage, nuclear condensation, and the activation of endonucleases. The chief effectors of the apoptotic cell death pathway are the caspase family of cysteine proteases. Caspases are synthesized as inactive precursors (procaspases) that must be cleaved at specific aspartate residues to generate the active (cleaved) subunits. Procaspase cleavage can occur by several mechanisms including proximity-induced autoprocessing or cleavage by other caspases, revealing a caspase cascade with upstream initiator caspases such as caspases-8, -9, and -10 and downstream effector caspases such as caspases -3, -6, and -7. The activity of these caspases is regulated by several families of both pro- and antiapoptotic cellular proteins (for reviews, see Refs.12 , 13).
In this study, in BPH tissues from patients treated for different periods with finasteride, we addressed the question as to whether the apoptotic process is caspase dependent. For this purpose, we immunolocalized and examined at different periods of finasteride treatment the expression and activation of two effector caspases (caspase-3, -6) and two initiator caspases (caspase-8, -9).
Subjects and Methods
A retrospective analysis was performed on patients treated or not with finasteride at the Department of Urology at Centre Hospitalier Lyon-Sud. According to the length of finasteride (5 mg/d) treatment, four groups of patients were determined: untreated (control) patients (n = 10), patients treated for 2–3 d (n = 7), patients treated for 6–8 d (n = 8), and patients treated for 27–32 d (n = 4). A total of 29 patients were included in this study. The mean age of the patients was 72 yr (range 58–93 yr). Characteristics of the patients in terms of age, plasma prostate-specific antigen (PSA) and prostate volume before finasteride treatment are presented in Table 1. The different patient groups are homogeneous because no statistical differences for the different parameters (age, PSA, prostate volume) among the groups were observed. Patients taking other medication known to possibly influence androgen levels were excluded. Endourethral resection was performed 2–3, 6–8,or 27–32 d after treatment. The fragments were either fixed in Bouin’s liquid or frozen in liquid nitrogen. All the cases were reviewed by a pathologist (M.Dec.), and the diagnosis of BPH was confirmed. For each case, a representative block of the BPH tissues was chosen for the immunohistochemical study. The ethics committee of the medical faculty and the state medical board agreed to these investigations, and informed consent was obtained for all patients.
Materials
Horseradish peroxidase-labeled antirabbit IgG and chemiluminescent kit were obtained from CovalAb (Lyon, France). Horseradish peroxidase-labeled antigoat IgG (SC-2020), rabbit polyclonal antibody raised against caspase-9, which reacts with both procaspase-9 (Mr 35,000) and cleaved caspase-9 (Mr 24,000), goat polyclonal antibody raised against caspase-8, which reacts with both procaspase-8 (Mr 55,000) and cleaved caspase-8 (Mr 25,000), and rabbit polyclonal antibody raised against caspase-3, which reacts with both procaspase-3 (Mr 30,000) and cleaved caspase-3 (Mr 17,000) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal antibody raised against cleaved caspase-3, which detects only the large fragment of activated caspase-3 (Mr 17,000), and rabbit polyclonal antibody raised against caspase-6, which detects procaspase-6 (Mr 34,000) and activated caspase-6 (Mr 12,000), were obtained from Ozyme (Saint Quentin en Yvelines, France). Mouse monoclonal keratin-903 antibody was obtained from Enzo Diagnostics, Inc. (Farmingdale, NY). The Envision+ kit (mouse or rabbit), the LSAB kit, Meyer’s hematoxylin, and Faramount were obtained from Dako (Trappes, France). Superfrost plus glass slides were obtained from Menzel-Glaser (Frelburg, Germany). Thymus terminal deoxynucleotidyl transferase, biotin deoxyuridine 5-triphosphate (dUTP), LC FastStart master SYBR green kit, and LC capillaries were obtained from Roche Molecular Biochemicals (Mannheim, Germany). Cobalt chloride, sodium cacodylate, extravidin peroxidase, diaminobenzidine (DAB), nickel chloride, and random primers were purchased from Sigma (Meylan, France). Finasteride (Proscar) was obtained from MSD (Whitehouse Station, NJ). Dithiothreitol, deoxynucleotide triphosphates, and Moloney murine leukemia virus reverse transcriptase were obtained from Invitrogen SARL (Cergy Pontoise, France).
Methods
Terminal deoxynucleotidyl transferase-mediated dUTP nick labeling (TUNEL). Paraffin sections (5 μm) of Bouin-fixed prostatic tissues were mounted onto Superfrost plus slides. The sections were deparaffinized and rehydrated (xylene 5 min, ethanol 100, 95, 70%, 30 sec each) and then washed in distilled water before beginning the TUNEL reaction. The slides were transferred to a plastic jar containing 0.01 M citrate buffer (pH 6) and microwave irradiated for 5 min (370 W) and then left 20 min at room temperature (RT). After a wash with phosphate buffer saline (PBSx 1), endogenous peroxidases were blocked for 5 min with H2O2 2%. Sections were washed three times with PBSx 1. The specimens were then incubated 60 min at 37 C in a moist chamber with TUNEL mix, which consisted of 0.3 U/μl calf thymus terminal deoxynucleotidyl transferase, 0.007 nmol/μl biotin dUTP, 1 mM cobalt chloride, 30 mM Tris (pH 7.2), and 140 mM sodium cacodylate. After washing (four PBS baths of 5 min each at RT) the sections were saturated 10 min with BSA 2% at RT. Sections were treated for 30 min at 37 C in moist chamber with a 1:20 dilution of Extravidin peroxidase. After three washes in PBS, detection was performed with DAB (1.25 mg), 25 μl nickel chloride 3%, 152 μl Tris-HCl 1 M (pH 7.5), completed with distilled water to 2 ml). Slides were mounted in Faramount.
SYBR green real-time PCR. Total RNAs were extracted from BPH tissues with TRIzol reagent (untreated patients, n = 7; patients treated with finasteride for 2–3 d, n = 4; for 6–8 d, n = 4; for 28–32 d, n = 6). cDNAs were obtained from 1 μg RNA incubated 1 h at 37 C in the presence of 50 μM random primers, 200 μM deoxynucleotide triphosphates, 100 mM dithiothreitol, and 200 U/μl of Moloney murine leukemia virus in a total volume of 20 μl.
In a first step, we assessed standards for caspase (-3 and -6) and ?-actin RNA quantitation by SYBR green PCR. The standards were assessed by scalar dilution of caspase or ?-actin cDNA. The samples were run in triplicate.
SYBR green real-time PCR assays were carried out in 20 μl PCR mixture volume consisting of 2 μl of LC FastStart SYBR green master mix, containing HotStarTaq DNA polymerase, 1 μl of 10 μM of each oligonucleotide primer, 4 μl of diluted RT mix (1/40) and 5 mM MgCl2.
Caspase-3 or -6 or ?-actin gene amplification was carried out as follows: initial activation of HotStarTaq DNA polymerase at 95 C for 480 sec; 45 cycles in three steps: 95 C for 15 sec, 55 C (caspase-3), 59 C (caspase-6), or 68 C (?-actin) for 6 sec, and 72 C for 12 sec. At the end of amplification cycles, melting temperature analysis was carried out by a slow increase in temperature (0.1 C/sec) up to 95 C. The primers used were: caspase-3, 5'-GAGCTGCCTGTAACTTG-3' and 5'-ACCTTTAGAACATTTCCACT-3'; caspase-6, 5'-ACTGGCTTGTTCAAAGG-3' and 5'-CAGCGTGTAAACGGAG-3’; and ?-actin (14). PCR products were checked by direct sequencing. Amplification, data acquisition, and analysis were carried out by LightCycler instrument (Roche Molecular Biochemicals) using LightCycler software (version 5.3.2, Roche). This software, coupled to the LightCycler instrument, determines the threshold cycle that represents the number of cycles in which the fluorescence intensity is significantly above the background fluorescence. The amounts of caspase (-3 or -6) relative to the amount of ?-actin in the same sample were analyzed by the relative quantification approach with the use of efficiency correction using Real Quant 1.0 software (Roche Molecular Biochemicals). The results are expressed as a normalized ratio.
Western blotting analysis. Proteins were prepared from tissue samples frozen in liquid nitrogen and then stored at –70 C until use. Prostatic tissues were homogenized in 200 μl ice-cold hypotonic buffer [25 mM Tris-HCl (pH 7.4), protease inhibitor cocktail] and sonicated (10 sec at 80). Protein concentration was determined by the Bradford assay. For Western blot, three patients for each experimental condition were used. The same antibodies were used for immunohistochemistry and Western blot analyses.
Proteins (100 μg) were resolved on 10% sodium dodecyl sulfate/polyacrylamide gels and electrophoretically transferred to nitrocellulose membranes using Tris 15 mM-glycine 120 mM buffer (pH 8.3) containing 20% methanol at a constant voltage of 100 V for 30 min. Under those conditions, the transfer of low-molecular-weight proteins is favored, i.e. transfer of activated caspases (which Mr ranges from 12,000 to 20,000). After transfer, the membranes were incubated in a blocking buffer [Tris-buffered saline (TBS) containing 5% fat-free dry milk and 0.1% Tween 20] 2 h at RT, rinsed three times with TBS/0.1% Tween 20 (3 x 10 min), and then incubated with the first antibody (in TBS containing 2% fat-free dry milk) overnight at 4 C. Anti-caspase-3 and anticleaved caspase-3 were diluted at 1/100, anticaspase-6 was diluted at 1/200, and anticaspase-8 and -9 were diluted at 1/100. The membranes were then rinsed with TBS/0.1% Tween 20 (3 x 10 min) and incubated with horseradish peroxidase-labeled antirabbit IgG (1/2000) or antigoat IgG (1/3000) in TBS containing 1% fat-free dry milk and 0.1% Tween 20 for 1 h at RT. The membranes were thoroughly washed with TBS/0.1% Tween 20 (3 x 10 min) and then with TBS. Bound antibodies were detected by the chemiluminescence kit and Biomax MR films (Sigma, Meylan, France). Protein loading was checked by reprobing the blot with a rabbit anti-actin IgG (1/500). The Biomax MR films were scanned on a Gel doc 2000 apparatus (BioRad Life Science Group, Marnes La Coquette, France), and quantitation was realized with Quantity one Software (Bio-Rad Life Science Group). The results are expressed as a ratio of caspase to actin.
Immunohistochemistry. Paraffin sections of Bouin-fixed prostatic tissues were sectioned at 5 μm. The sections were mounted on Superfrost plus glass slides, deparaffinized, hydrated, treated 20 min at 93–98 C in citric buffer [0.01 M (pH 6)], rinsed in osmosed water (2 x 5 min), and washed (2 x 5 min) in TBS (pH 7.8). For anticaspase-3 (pro and cleaved), anticleaved caspase-3, anticaspase-6, and anticaspase-9 antibodies, the Envision plus kit was used. For the anticaspase-8 antibody, the LSAB kit was used. The first steps were similar for both kits. Briefly, endogenous peroxidases were blocked with 3% H2O2 for 15 min and then washed three times in TBS. The sections were then incubated overnight at 4 C with the primary antibody diluted in antibody diluent (anticaspase-3, 1/100; anticleaved caspase-3, 1/50; anticaspase-6, anticaspase-9, and anticaspase-8, 1/100). The sections were then washed (2 x 5 min) in TBS. For the Envision plus kit, the sections were incubated 30 min at 37 C in presence of the secondary antibody peroxidase-conjugated. The sections were then rinsed (2 x 5 min) in TBS, incubated 10 min at RT with 3-amino-9-ethylcarbazole or DAB, which generated a red color at the site of peroxidase activity and rinsed (2 x 5 min) in osmosed water. For the LSAB kit, after incubation with the primary antibody, the section were incubated with the biotinylated secondary antibody, and after washes, a peroxidase streptavidin complex was applied. DAB or 3-amino-9-ethylcarbazole was used as a peroxidase chromogen. For double-staining experiments, the slides were first incubated with the keratin-903 primary antibody (1/200). After washes, the slides were incubated with an antimouse secondary antibody peroxidase-conjugated (mouse Envision plus kit) and revealed with DAB. After washing, the slides were incubated with the anticleaved caspase-3 antibody (1/50). The antigen-antibody complexes were revealed with the LSAB kit with a streptavidin phosphatase conjugate and revealed with fast red. For the different experiments, sections were counterstained with Meyer’s hematoxylin. Finally, sections were mounted in Faramount. As a negative control, the primary antibody was replaced by the antibody diluent. Immunohistochemistry was performed on a slide from each patient. For analysis of immunohistochemical staining, the intensity was rated as none, weak, moderate, or intense for each slide. Specimens were considered immunopositive when 1% or more of the BPH tissue had clear evidence of immunostaining. The immunostaining was evaluated by two independent observers in the laboratory, blinded as to the treatment status. Similar results were obtained for the different patients; a representative immunohistochemistry is presented.
Data analysis. Depending on the availability of BPH tissues, the number of patients studied in the different groups is as follows: four to 10 patients per condition for the immunohistochemical experiments, three patients per condition for the Western blotting analyses, and four to seven per condition for the real-time PCR experiments. For statistical analysis one-way ANOVA was performed to determine whether there were differences among all groups (P < 0.05), and then the Bonferroni posttest was performed to determine the significance of the differences between the pair of groups. P < 0.05 was considered significant. The statistical tests were performed on StatView software (version 5.0, SAS Institute Inc., Cary, NC).
Results
Time dependency of the apoptotic process in BPH tissues on finasteride administration
Very few TUNEL-positive (apoptotic) cells were observed in BPH tissues from untreated patients (Fig. 1A). In contrast, a higher amount of TUNEL-positive cells were visualized in epithelial secretory cells of prostatic tissues from patients affected by BPH and treated during 2–3 or 6–8 d with finasteride (Fig. 1B). A very limited number of apoptotic cells was observed in stromal cells (Fig. 1, A and B).
Consistently, the evaluation of the number of TUNEL-positive cells in BPH tissues indicated an increase in apoptotic cells after 2–3 d of treatment and a more dramatic increase after 6–8 d (250-fold, P < 0.0001). However, after 27–32 d of finasteride treatment, the apoptotic cell number decreased and was back to control values (Fig. 1C).
To further characterize the apoptotic cell death process in prostatic epithelial cells, four caspases, two effector caspases (caspase-3 and -6) and two initiator caspases (caspase-9 and –8), were studied in terms of proteins in BPH tissues from untreated or finasteride-treated patients.
Immunolocalization of caspases-3, -6, -9, and -8 in BPH tissues after finasteride treatment
The immunohistochemical experiments were performed in BPH tissues from each patient, treated or not treated with finasteride. A representative experiment is presented for each different caspase. For the immunolocalization of caspase-3, two different antibodies were used. The first one detected both pro- and cleaved caspase-3 (Fig. 2, A–D), whereas the second one detected exclusively cleaved (activated) caspase-3 (Fig. 2, E–H). In tissues from untreated patients, caspase-3 was mainly detected in basal and secretory epithelial cells and to a lesser extent in stromal cells (Fig. 2A). After finasteride treatment, caspase-3 immunolocalization in BPH tissues was not affected (Fig. 2), although an apparent increase in caspase-3 immunostaining intensity, depending on the different times of treatment including 2–3 d (Fig. 2B) and 6–8 d (Fig. 2C), could be observed. Interestingly, cleaved caspase-3 immunostaining was either not detected or detected at very low levels in BPH tissues from untreated patients (Fig. 2E), whereas, in contrast, after treatment with finasteride for 2–3 and 6–8 d, its staining increased and appeared exclusively in the epithelial secretory cells (Fig. 2, F and G). Then 27–32 d after finasteride treatment, the cleaved caspase-3 staining intensity was reduced and appeared to be similar to that observed in tissues from untreated patients (Fig. 2H). To further evaluate whether activated caspase-3 was expressed in the secretory or basal epithelial cells of prostate, we used an antibody raised against keratin-903, known to be expressed exclusively in the basal epithelial cells. Keratin-903 staining was observed in the basal epithelial cells of BPH tissues from both untreated patients and patients treated with finasteride for 2–3, 6–8, and 28–32 d (Fig. 2, I–L). Activated caspase-3 staining was detected (exclusively) in secretory epithelial cells but not in basal cells of BPH tissues from patients treated with finasteride for 2–3 and 6–8 d (Fig. 2, J and K). In BPH tissues from patients treated with finasteride for 28–32 d, prostatic secretory epithelial cells remained detectable, although activated caspase-3 staining was no longer observed (Fig. 2L). Such an observation suggests that the decrease in activated caspase-3 staining in tissues from patients treated with finasteride for 28–32 d was probably not related to epithelial cell loss.
For the immunolocalization of caspase-6, the antibody used recognized both pro- and cleaved caspase-6. In BPH tissues from both control untreated and treated patients, caspase-6 was immunolocalized to secretory and basal cells and to a lesser extent to stromal cells (Fig. 3A). However, an apparent increase in the intensity of caspase-6 staining in the epithelial secretory cells could be observed in tissues from patients treated for 2–3 and 6–8 d with finasteride (Fig. 3, B and C). After 27–32 d of treatment, the caspase-6 staining intensity appeared to be reduced and was back to the staining intensity observed in tissues from untreated patients (Fig. 3D).
Antibodies used for the detection of initiator caspases (caspase-9 and -8) were designed to recognize both pro- and cleaved caspases. Caspase-8 was immunodetected in the epithelial secretory, basal, and the stromal cells from the untreated patients (Fig. 3E). The intensity of caspase-8 immunostaining appeared not to be affected by finasteride treatment in BPH tissues (Fig. 3, F–H). In BPH tissues, caspase-9 was immunoexpressed in the secretory epithelial and stromal cells (Fig. 3I). In contrast to caspase-8, the intensity of caspase-9 immunostaining appeared to be higher in BPH tissues from patients after 2–3 or 6–8 d of treatment (Fig. 3, J and K), whereas it was reduced and was similar to that observed in tissues from patients treated with finasteride for 27–32 d (Fig. 3L).
To strengthen the data obtained from the immunohistochemical experiments, we further carried out quantitative studies in BPH tissues by Western blotting analyses on caspase protein levels and real-time quantitative PCR on caspase mRNA levels.
Effects of finasteride treatment on caspase-3, -6, -8, and -9 protein levels in BPH tissues
The same antibodies used for the previous immunohistochemical studies were also used for Western blotting experiments. The Western blotting experimental conditions were optimized to detect the active-cleaved caspases (see Materials and Methods). Activated caspases [caspase-3 (Mr 17,000) and -6 (Mr 12,000)] protein levels were low in untreated patients, whereas their levels were significantly (P < 0.0003) increased after a 6–8 d of finasteride treatment (Fig. 4, A and B). Caspase-8 was not detected in terms of cleaved caspase in BPH tissues from patients treated or not with finasteride (data not shown). The cleaved caspase-9 protein (Mr 20,000) was detected at low levels in untreated patient tissues, but such levels were significantly (P < 0.0113) increased in tissues from patients treated during 2–3 and to a lesser extent during 6–8 d of treatment with finasteride (Fig. 4C).
Effects of finasteride treatment on caspase-3 and -6 mRNA levels in BPH tissues
Caspase-3 and -6 mRNA levels in BPH tissues from untreated patients or patients treated for 2–3, 6–8, or 27–32 d with finasteride were determined by using real-time quantitative RT-PCR assays. No significant changes in caspase-3 or -6 mRNA levels were observed after finasteride treatment (Fig. 5, A and B).
Discussion
Among the therapies used against BPH (at least with a prostate weight >40 g), the treatment with a 5-reductase inhibitor such as finasteride has been reported to be effective in improving both clinical symptoms and urinary flow rates as well as in reducing prostate size. Although it has been shown that BPH treated with finasteride exhibits a progressive decrease in prostatic cell size and function during the first several months of treatment (11), the molecular mechanisms underlying finasteride action remain poorly understood.
In the present study, finasteride was shown to trigger apoptosis in BPH tissues in the first week of treatment. Indeed, apoptotic epithelial prostatic cells were identified 2–3 d after finasteride treatment with the highest level of apoptotic cell number evidenced after 6–8 d of treatment. In contrast, after 27–32 d of treatment with the inhibitor, the number of apoptotic epithelial prostatic cells was back to levels comparable with those observed in tissues from untreated patients. Such observations are consistent with those reported by Rittmaster et al. (11), although these authors have used a long-term protocol (6–18 and 23–73 d and 3–48 wk of treatment), compared with our own protocol. Indeed, the peak of apoptosis after finasteride treatment was observed after 6–8 d (our study) or 6–18 d (11). Moreover, the apoptotic process was detected in the prostatic epithelial cells but not in stromal cells of BPH tissues treated with finasteride (the present study and Ref.11). Our present data suggest that the effects of finasteride treatment in the cell death process, at least evaluated through the TUNEL approach, appear to be transient. Indeed, the death process was observed after 2–3 and 6–8 d but not after 27–32 d of treatment. Second, these data confirm that the death process affects mainly epithelial cells but not stromal cells. This observation could explain that, although finasteride is effective in reducing prostate volume, such a decrease was only of 20–30% of the initial volume because it may primarily concern the epithelial cell component (4, 10, 15, 16).
To further characterize the molecular mechanisms underlying the finasteride-induced apoptotic process, we studied the caspase proteins that are key factors in the cell death machinery (for a review, see Ref.13). In this context, we characterized the expression of pro- and especially cleaved caspases in the apoptosis induced by finasteride in BPH tissues. The effector caspase-3 and -6 were shown to be expressed in secretory and basal epithelial cells and to a lesser extent in the stromal cells. Indeed, caspase-3 was immunodetected in basal and secretory epithelial cells by an antibody recognizing both pro- and activated caspase-3 (17). One of the original contributions of the present report is the demonstration that finasteride treatment induces the activation of caspases-3 and -6 during the apoptotic cell death process in BPH tissues. Indeed, an apparent increase in the immunostaining of the caspases-3 and -6 was observed when patients were treated during 2–3 or 6–8 d with the inhibitor. More specifically, activated caspase-3, which was shown to be immunolocalized exclusively in secretory but not in basal epithelial cells of BPH tissues, exhibited an increased immunostaining 2–3 and 6–8 d after treatment with finasteride. In contrast, activated caspase-3 was not detected (or very weakly) in untreated tissues and after 27–32 d of treatment with finasteride, specifically, activated caspase-3 was no longer detected (or very weakly) in the secretory epithelial cells of the prostate. The dramatic decrease of the activated caspase-3 staining was not related to a loss of epithelial secretory cells because these cells remained detectable. Furthermore, these results were confirmed by Western blotting experiments showing an increase in activated caspase-3 and -6 protein levels in BPH by 2–3 d and at higher levels by 6–8 d in treated patients. The changes in caspase-3 and -6 levels were related to alterations of protein but not of mRNA levels. Interestingly, the activation of caspase-3 and -6 was kinetically correlated with the apoptotic process in epithelial prostatic cells because the TUNEL approach shows a peak of cell death by 6–8 d in BPH patients treated with finasteride. The absence of activated caspases in stromal prostatic cells suggests that these cells probably do not use the caspase-3 pathway to trigger cell death. Some authors suggested that stromal cells of BPH may use the caspase-7 pathway to trigger apoptosis (18).
At least two pathways, the death receptor and the mitochondrial pathways, are known to activate the effector caspases via the initiator caspases. The death receptor pathway triggers the activation of the initiator caspase-8 or -10, whereas the mitochondrial pathway triggers the activation of initiator caspase-9 (19). We examined here the expression of initiator caspases belonging to each pathway, i.e. caspase-8 and -9. Caspase-8, which was detected in human prostate cancer cell lines LNCaP (20), exhibited a low immunostaining intensity in both stromal and epithelial cells in our experiments. Moreover, finasteride treatment had no effect on the caspase-8 protein expression, suggesting that finasteride-induced apoptosis in prostatic epithelial cells probably did not result from the activation of the caspase-8 pathway. However, it cannot be excluded that caspase-8 is at play in prostate cancer cell line apoptosis. Indeed, in LNCaP cell lines, TNF, and ionizing radiations induce pro- and activated caspase-8 expressions, and an addition of androgens inhibits this caspase-8 activation (21). Based on our findings, it is quite possible that by contrast to effector caspase-3 and-6, caspase-8 is not targeted by androgen action in BPH tissues. This is consistent with our recent data showing that, whereas rat castration induces, in ventral prostate, effector caspase-3 and -6 activation, it fails to affect caspase-8 expression or activation (22).
With regard to the mitochondrial pathway, we identified caspase-9 in epithelial and stromal cells in untreated BPH tissues. The immunostaining intensity of caspase-9 appeared to increase by 2–3 d and to a lesser extent 6–8 d after finasteride treatment. This observation was confirmed by Western blotting experiments, showing a maximal increase in cleaved caspase-9 protein levels after 2–3 d of treatment, suggesting that executor caspases-3 and -6 may be activated in the mitochondrial pathway. Indeed, maximal levels of activated caspase-9 were observed by 2–3 d before the peak of activated caspases-3 and -6 (observed by 6–8 d of finasteride treatment). It is of interest to mention that the possible involvement of the mitochondrial pathway during the apoptotic process in prostatic cells was also previously reported in prostate cancer cell lines. Indeed, procaspase-9 expression was detected in LNCaP cell lines (23), and withdrawal of androgens (castration) was shown to activate caspase-9 in rat prostate epithelial cells (24).
Together, finasteride triggers apoptosis exclusively in epithelial secretory cells, and its action on the apoptotic process was transient because it was limited to a short period (<27–32 d). No TUNEL-positive cells were observed after 27–32 (our present study) or 23–73 d (11) of treatment with finasteride, and effector caspase-3 was no more activated after 27–32 d of treatment with finasteride, although epithelial cells were still present because they were detectable (our present study). The molecular mechanisms underlying this arrest in the cell death process after 1 month of treatment with finasteride remain unknown. However, despite an arrest of the finasteride-dependent apoptotic process after 1 month, the inhibitor has been shown to sustain a reduced prostate volume after 1–6 yr of therapy (10, 16, 25, 26). Whether mechanisms/processes other than apoptosis could underline the BPH tissue atrophy in the long-term treatment remains to be answered.
In summary, by using BPH tissues from patients treated with finasteride during, 2–3, 6–8, or 27–32 d, we show that the antiandrogen induces apoptosis in prostatic epithelial cells within 2–3 to 6–8 d of treatment but not later (27–32 d). Such an apoptotic process is caspase dependent because it appears to result from activation of effector caspases-3 and -6, probably involving the mitochondrial pathway as evidenced by the activation of caspase 9.
Acknowledgments
We are grateful to Dr. A. Florin-McLeer for critical reading of the manuscript.
Footnotes
This work was supported by Institut National de la Santé et de la Recherche Médicale; Ministère de l’Enseignement Supérieur et de la Recherche; and la Ligue Contre le Cancer de l’Ardèche, France.
First Published Online October 26, 2004
1 A.B. and A.R. contributed equally to the work in this manuscript.
Abbreviations: BPH, Benign prostatic hyperplasia; DAB, diaminobenzidine; DHT, dihydrotestosterone; dUTP, deoxyuridine 5-triphosphate; PSA, prostate-specific antigen; RT, room temperature; TBS, Tris-buffered saline; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick labeling.
Received April 22, 2004.
Accepted October 19, 2004.
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