Background

Apoptosis is a mode of cell death that participates in the kidney physiologic remodeling processes and is thought to contribute to cell loss and kidney structural damage in chronic renal diseases. Gender is one factor which contributes to accelerated nephron loss, with progression more rapid in men than in women in diabetic and nondiabetic chronic renal diseases. Mechanisms by which androgens may cause higher rate of progression of chronic renal diseases in men are poorly explored.

Methods

In this study, to investigate the role of androgens on apoptotic damage and its associated mechanisms, we examined the effects of testosterone (T) (0.1 nmol/L to 1 mol/L) on apoptosis, and apoptosis-related proteins in a proximal human tubule cell line (HK-2 cells). Additional experiments were performed in primary cultures of proximal tubular epithelial cells (PTECs). Cells were grown to subconfluence in normal growth medium, and apoptotic damage was induced by serum deprivation for 24 to 48 hours. Cycloheximide, flutamide (a T-receptor antagonist), 17- estradiol, or caspase inhibitors were added to cultures that were successively processed for terminal deoxynucleotidyl transferase-mediated uridine triphosphate nick end-labeling (TUNEL) analysis, annexin V/propidium iodide staining, immunofluorescence, or immunoblots to identify effects and apoptotic pathways that could be modulating cell survival.

Results

Both morphologic analysis by annexin V/propidium iodide staining and TUNEL showed that physiologic T levels (1 to 10 nmol/L) induced a significant increase in apoptosis both in HK-2 cells and PTECs. In both types of cell lines pretreatment with the androgen receptor antagonist flutamide prevented the T-induced apoptosis. T-induced apoptosis was enhanced by treatment with cycloheximide and prevented by 17-estradiol. Fas, Fas ligand (FasL), and Fas-associating death domain containing protein (FADD) were clearly up-regulated within 48 hours of T treatment in HK-2 cells. Also, T significantly increased the expression of Bax protein (P < 0.01 vs. control) (an effect which was blocked by flutamide), and decreased the expression of Bcl-2. Western blot analysis showed that caspase-3 was activated. Moreover, cleavage into an 85-kD poly(ADP-ribose) polymerase-1 (PARP-1) terminal breakdown product was detectable. The changes in cellular morphology induced by T at 48 hours were no longer observed after the addition of caspase-8, caspase-9, and caspase-3 inhibitors to the culture medium.

Conclusion

These results indicate that T increases the permissiveness of proximal tubule kidney cells to apoptotic effects by triggering an apoptotic pathway involving caspase activation, Fas up-regulation, and FasL expression, thus potentially interacting with mechanisms of cell loss which have been already shown to be activated in chronic renal diseases. This is consistent with a role for T in promoting renal injury in men.

METHODS

Reagents

Media and additives for culture of HK-2 cells, as well as annexin fluoroscein isothiocyanate (FITC), propidium iodide, 3,3-diaminobenzidine (DAB), testosterone, flutamide, 17-estradiol, phenazine methosulfate, p-iodonitrotetrazolium violet, -NAD, L-(+)-lactic acid, cycloheximide, acrylamide, and other reagents for sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) were purchased from Sigma Chemical Company (St. Louis, MO, USA). Caspase inhibitors were purchased from R&D Systems Europe, Ltd. (Oxon, UK). Coomassie protein assay reagent was obtained from Pierce (Rockford, IL, USA). Tissue culture plates and dishes were from Corning (New York, NY, USA). Antihuman Bcl-2 monoclonal antibody, antihuman Bax polyclonal antobody, antihuman Fas and FasL antibodies were obtained from Upstate Biotechnology (Lake Placid, NY, USA). Antihuman-cleaved poly (ADP-ribose) polymerase-1 (PARP-1) polyclonal antibody was purchased from Alexis Biochemicals (Lausen, Switzerland). Antihuman FADD (monoclonal antibody) and antiactive caspase-3 (p17) monoclonal antibody were purchased from BioVision Research Products (Mountain View, CA, USA).

The biotin-streptavidin-amplified detection system was obtained from Biogenex (Biogenex Laboratories, San Ramon, CA, USA) and Eukitt from O Kindler GmbH and Co. (Freiburg, Germany). Terminal deoxynucleotidyl transferase (TdT), TdT buffer, and biotynilated dUTP were obtained from Boehringer Mannheim (Mannheim, Germany). Hybond-c-nitrocellulose membrane, enhanced chemiluminescent reaction ECL, and Hyperfilm-ECL were obtained from Amersham Pharmacia Biotech (Arlington Heights, IL, USA).

Cell culture

HK-2 cells, an immortalized proximal tubular epithelial cell line from normal adult human male kidney, were obtained from American Tissue Culture Collection (ATCC, Manassas, VA, USA). Primary human proximal tubular epithelial cells (PTECs) were cultured according to the method of Detrisac et al¹⁸. Portions of renal cortex not involved by disease were obtained from male kidneys removed by surgery for renal cell carcinoma. Small fragments of renal tissue were placed on a matrix of type I bovine collagen and fetal bovine serum (FBS). Outgrowth of PTECs was confirmed by their morphologic appearance, enzyme cytochemistry (reactivity for alkaline phosphatase, gamma-glutamyltranspeptidase, acid phosphatase, and leucine aminopeptidase), and immunofluorescence. PTECs were used between the third and fifth passage. Cells were grown in phenol red-free Dulbecco's modified Eagle's medium (DMEM)/F12 medium supplemented with 5% (v/v) FBS, 100 U/mL penicillin-streptomycin, 2 mmol L-glutamine, 5 g/mL insulin, 5 g/mL transferrin, 5 ng/mL sodium selenite, 5 pg/mL T3, 5 ng/mL hydrocortisone, 5 pg/mL PGE1, and 10 ng/mL epidermal growth factor. Cells were grown at 37°C in a humidified 5% CO₂ condition.

Cell treatments

Cells were grown to subconfluence in normal growth medium, and apoptotic damage was induced by additives and serum deprivation for 24 to 48 hours in the presence of T (0.1 nmol/L to 1 mol/L). Control cells (C) were not treated with hormone. In order to determine if T is directly involved in apoptosis, the antiandrogen flutamide (100 nmol/L) was added 4 hours earlier than 10 nmol/L testosterone. In separate settings of experiments, cycloheximide (10 g/mL), a combination of T and cycloheximide, or buffer control, respectively, was used. In other experiments, 17-estradiol (10 nmol/L) was used in combination with T. To determine a role of caspases in 10 nmol/L T-induced cell death, HK-2 cells were preincubated for 30 minutes with 50 mol/L caspase inhibitors. Caspase-8 inhibitor II (Z-IEDT-FMK), caspase-9 inhibitor I (Z-LEHD-FMK), and caspase-3 inhibitor I (DEVD-CHO) were used.

Cell viability

Experiments were carried out by adding T at scalar doses to 10⁵ cells cultured in 12-well plates. Nontreated cells were used as control. After 24 and 48 hours of incubation, cells were counted using a NeuBauer chamber (Corning Costar Co., Cambridge, MA, USA). Cell viability was assessed by Trypan blue. For each treatment group the number of adherent cells at t = 0 served as baseline value (100%), and was used to express the percentage of adherent cells.

Lactate dehydrogenase release

The amount of lactate dehydrogenase (LDH) activity released into the medium was measured to assess the leakage of normal components of the cytoplasm into the medium. For LDH assay, supernatant aliquots were transferred to the corresponding wells of a 96-well plate. Then 100 L of substrate containing L-(+)-lactic acid, -NAD, p-iodonitrotetrazolium violet, phenazine methosulfate, and Tris buffer were added at each well and incubated at 37°C until reaching a plateau. The absorbance was calculated at 540 nm using a scanning multiwell spectrophotometer.

Annexin V-FITC/propidium iodide (PI) staining

In multiple experiments, HK-2 cells were stained with annexin V/propidium iodide and observed under a fluorescence microscope. The assay is based on the ability of annexin V (green fluorescence) to bind to the phosphatidylserine exposed on the surface of cells undergoing apoptosis and the capacity of propidium iodide (red fluorescence) to enter cells that have lost their membrane integrity. HK-2 cells were grown on chamber slides, washed with Dulbecco's phosphate buffer solution (DPBS), and incubated in the dark for 5 minutes at room temperature with 200 L of 1 binding buffer, 1 L of FITC-labeled recombinant annexin V, and 1 L of propidium iodide. Cells were observed and counted (300 cells for each condition and experiment) under a fluorescence microscope using dual filter set for FITC and rhodamine. Cells that lost membrane integrity showed red staining throughout the nucleus and a halo of green staining on the cell surface. To evaluate apoptotic phenomena, we considered the percentage of cells annexin V–positive/propidium iodide–negative.

Terminal deoxynucleotidyl transferase-mediated uridine triphosphate nick end-labeling (TUNEL)

HK-2 cells grown on chamber slides to subconfluence were incubated for 48 hours with or without T (10 nmol/L). After a five-minute incubation in 2% paraformaldehyde, cells were fixed in 2:1 vol/vol ethanol:acetic acid at room temperature for 10 minutes. After three washes with phosphate-buffered saline (PBS), cells were incubated with 100 U/mL TdT, 0.5 g/mL biotynilated uridine triphosphate in 140 mmol/L potassium cacodylate, 125 mmol/L Tris-HCl, 2.5 mmol/L cobalt chloride, pH 6.6, for 1 hour at 37°C in a humidified chamber. After stopping the reaction by transferring the slides to tris-borate (TB) buffer (300 mmol/L sodium chloride, 30 mmol/L sodium citrate) and washing in PBS, cells were incubated for 30 minutes at room temperature with extra-avidin-peroxidase diluted 1:20 in PBS, and then incubated with the peroxidase-substrate solution (0.04% 3,3-diaminobenzidine in 50 mmol/L Tris-HCl buffer containing 0.03% hydrogen peroxide) for 5 minutes. After rinsing with PBS, sections were counterstained with hematoxylin and examined by light microscopy.

Cell immunofluorescence and immunostaining

The expression of Fas, FasL, FADD, and p17 fragment of the active caspase-3 protein were initially documented by immunofluorescence. HK-2 cells were grown on chamber slides to subconfluence and incubated for 48 hours in the presence or absence of 10 nmol/L T. Cells were then washed with cold PBS and fixed in cold methanol for 5 minutes. After a 30-minute incubation with anti-FAS (polyclonal Ab 1:40 in PBS), anti-FASL (polyclonal Ab 1:40 in PBS), anti-FADD (monoclonal Ab, 20 mg/mL), or antiactive caspase (polyclonal Ab, 20 mg/mL) antibodies at room temperature, cells were washed extensively with PBS and exposed to biotinylated conjugated secondary antibody for 30 minutes. After being washed, cells were incubated with FITC-streptavidin for 30 minutes. Slides were observed under a fluorescence microscope. The expression of cleaved PARP-1, Bax, and Bcl-2 proteins was documented by immunostaining. Both attached and floating cells were harvested by trypsin and incubated on poly-L-lysine coated glass slides for 40 minutes at 4°C. Then, the spots were air-dried and fixed in cold acetone for 30 seconds and stored at -20°C until the immunodetermination of cleaved PARP-1, Bax, and Bcl-2. After rehydration in PBS, spots were incubated with the antihuman cleaved PARP-1 polyclonal antibody (dilution 1 g/mL in PBS), anti-Bax polyclonal antibody (1:200 in PBS), and anti-Bcl-2 monoclonal antibody (1:200 in PBS) at room temperature. The second and third steps were performed using the improved biotin-streptavidin-amplified detection system. Briefly, cells were incubated with the secondary antibody (biotinylated IgG; 1:100 in PBS) for 20 minutes. After several washes with PBS, cells were incubated with the concentrated enzyme label (biotin-streptavidin-peroxidase) for 20 minutes at room temperature. Peroxidase was developed with 0.04% 3,3-diaminobenzidine in 50 mmol/L Tris-HCl buffer containing 0.03% hydrogen peroxide for 15 minutes. After rinsing with PBS, slides were counterstained with hematoxylin, coverslipped with Eukitt, and examined by light microscopy.

Image analysis

Image analysis was performed by the Leica Q500 MC Image Analysis System (Leica, Cambridge, UK). For each sample 300 cells were randomly analyzed and the optical density of the signals was quantitated by a PC computer. The video image was generated by a CCD Camera (Leica, Cambridge, UK) connected through a frame grabber to a PC computer. Single images were digitized for image analysis at 256 gray levels. Imported data were analyzed quantitatively by Q500MC Software-Qwin (Leica). The single cells were randomly selected by the operators using the cursor, and then positive areas automatically estimated. Constant optical threshold and filter combination were used^19,20.

Western blot analysis

The cell layers (including the floating cells) were washed twice with cold DPBS and cold lysis buffer [20 mmol/L HEPES, 150 mmol/L NaCl, 10% (v/v) glycerol, 0.5% (v/v) NP-40, 1 mmol/L EDTA, 2.5 mmol/L DTT, 10 g/L aprotinin, leupeptin, pepstatin A, 1mmol/L PMSF, and Na₃VO₄]. Cells lysates were incubated for 1 hour at 4°C, and the insoluble material was removed by centrifugation at 13,000 rpm for 20 minutes at 4°C. Protein concentration was determined by using the Coomassie protein reagent. Protein (30 or 50 g) was loaded on 10% to 15% SDS-PAGE under reducing or nonreducing conditions after being heated for 3 minutes at 100°C. The proteins were electrotransferred to a nitrocellulose membrane. Blots were blocked for 1 hour at room temperature in PBS 5% milk. The membrane was incubated overnight at 4°C with antihuman BAX, antihuman BCL-2, antihuman Fas, antihuman FasL, anti-FADD, antiactive caspase-3, (p17 fragment), anti-cleaved PARP-1 (p85 fragment), followed by three washes (total 30 minutes) with PBS 0.05% (v/v) Tween 20. The membranes were incubated with horseradish peroxidase-conjugated secondary antibodies for 1 hour at room temperature. Immunoblots were developed with the ECL Western blotting detection system.

Statistical analysis

Statistical analysis was performed with the InStat software package version 2.01 (Graph Pad, San Diego, CA, USA). The one-way analysis of variance (ANOVA) and the Tukey-Kramer multiple comparison test were used to test the significance of differences. Results are expressed as mean SD and are the expression of at least three experiments, with two wells for each experiment. Differences were considered statistically significant if P < 0.05.

RESULTS

Cell viability

During the course of the experiments, the majority of cells remained viable and attached to culture dishes in control cultures over a 48-hour period (88 6.5% of HK-2 cells and 93 6.4% of primary PTECs were viable, respectively, at 48 hours) Figure 1. To examine the apoptotic effects of androgens, cells were treated with different concentrations of T (0.1 nmol to 1 mol/L). When T was used at the lowest physiologic doses (0.1 to 10 nmol/L), 83 8% and 90.5 9% of HK-2 cells and PTECs, respectively, were viable at 24 hours (P > 0.05 vs. untreated cells). These percentages did not significantly change at 48 hours (80 2% and 80 11%, respectively, for HK-2 cells and PTECs) (Figure 1a and b). With 1 mol/L T (a concentration which is greater than that commonly found circulating in vivo in man)²¹, cell viability was significantly decreased at 48 hours (62 5% of HK-2 cells; 76 5% of PTECs viable).

Figure 1.

Effects of increasing doses of testosterone (T) on proximal human tubule cell line (HK-2) (A) and proximal tubular epithelial cells (PTEC) (B) viability at 24 () and 48 () hours. Cell viability was assessed by counting the number of adherent and floating cells. For each treatment group the number of cells at t = 0 served as baseline value 100% and was used to express the percentage of living cells. *P < 0.001 vs. control (C); ^#P < 0.05 vs. C. Effects of 10 nmol/L and 1 mol/L of T on HK-2 (C) and PTECs (D). LDH release at 48 hours. Results are expressed as mean SD and are representative of 3 separate experiments. Each one was carried out in triplicate. The differences were not statistically significant.

Full figure and legend (19K)

There were no significant changes in LDH release (Figure 1c and d) in untreated cells at 48 hours. Similarly, the addition of T did not cause cell injury, as represented by LDH release.

Effects of T on apoptosis in renal tubular cells

Cultured HK-2 cells and primary PTECs were incubated with different concentrations of T for 24 and 48 hours in multiple experiments and subjected to apoptosis assay(s). Treatment with T induced several apoptotic features both in HK-2 cells (Figure 2a – d) and PTECs. Features of the T-treated cells stained by TUNEL observed by light microscopy included rounding and detachment of the cells with fragmentation and shrinking of nuclei. Apoptotic cells showed chromatin condensation and fragmentation of nuclear material. The treatment with T also increased the percentage of annexin V/propidium iodide–positive cells, as shown by halo of green staining on cell surface Figure 2b. During the course of these experiments, only 3.60 1.93% of the untreated HK-2 cells was annexin V–positive at 24 hours (P = NS vs. t = 0), and this percentage increased slightly (9.7 3.01%) at 48 hours. T induced a progressive rise in apoptotic HK-2 cells Figure 3a, with a 2- to 3-fold increase being observed at 24 hours (12.3 2.5% and 14.6 2.46% for T 0.1 to 10 nmol/L, P < 0.05 to 0.01 vs. control). Analogous findings were observed at 48 hours Figure 3a. A similar trend was observed when cells were grown for 48 hours at different concentrations of T in 0.5% charcoal-stripped FBS (data not shown). For both T 10 nmol/L and 1 mol/L the apoptotic index was significantly higher than in experiments performed at 24 hours (P < 0.05 to 0.01).

Figure 2.

Apoptosis in proximal human tubule cell line (HK-2) cultures stained with annexin V (green fluorescence) and propidium iodide (red fluorescence). Control HK-2 (A), HK-2 treated with 10 nmol/L testosterone (T) for 48 hours (400) (B). DNA strand breaks as assessed by terminal deoxynucleotidyl transferase-mediated uridine triphosphate nick end-labeling (TUNEL) staining of HK-2. Cells were exposed or not exposed to 10 nmol/L T for 48 hours and subjected to TUNEL staining. Control HK-2 (C), HK-2 exposed to 10 nmol/L T (400) (D).

Full figure and legend (81K)

Figure 3.

Apoptosis detection by annexin V/propidium iodide. Subconfluent proximal human tubule cell line (HK-2) (A) and proximal tubular epithelial cells (PTECs) (B) were growth factor– and serum-deprived for 24 and 48 hours in presence of testosterone (T) (0.1 nmol/L to 1 mol/L). Cells were stained with annexin V/propidium iodide and examined under fluorescence microscope. Apoptotic cells were expressed as percent of total cells counted (350 cells for each condition). Data shown as mean SD. Each experiment was performed in triplicate. *P < 0.05; **P < 0.01; ***P < 0.001 vs. untreated cells (control); ^#P < 0.05 vs. 10 nmol/L 24 hours; ^##P < 0.01 vs. 1 mol/L 24 hours.

Full figure and legend (27K)

Similar proapoptotic effects of T were obtained in primary PTECs Figure 3b. At 24 hours, 4.3 2% of the untreated cells was annexin V–positive, and this percentage rose to 10.7 2.4% at 48 hours (P = NS vs. t = 24 hours). When primary PTECs were incubated with scalar doses of T, we observed a 3-fold increase in the apoptotic index at 24 hours (13.3 2.3% and 12 1% for 1 to 10 nmol/L T; P < 0.05 vs. C), which rose further at 48 hours (18 1.8% and 28.5 6.6% for T 1 nmol/L and 1 mol/L, respectively; P < 0.05 to 0.001). Similar to what was observed with the immortalized line, the apoptotic index at 48 hours was significantly higher than in experiments performed at 24 hours (P < 0.05 to 0.01).

Cycloheximide strongly potentiated the apoptotic effect induced by T. Apoptotic cell death involved approximately 15% of HK-2 cells after 48 hours, and this figure raised to 40% when cycloheximide was added to T Figure 4.

Figure 4.

Apoptosis detection by annexin V/propidium iodide. Subconfluent proximal human tubule cell line (HK-2) were growth factor– and serum-deprived for 24 hours in the presence or absence of testosterone (T) (10 nmol/L) and cycloeximide (10 g/mL) (CLX). *P < 0.05 vs. control (C); **P < 0.001 vs. T; ^#P < 0.001 vs. T; P < 0.001 vs. CLX.

Full figure and legend (7K)

Proapoptotic effects of T were abolished by the administration of the androgen-receptor antagonist flutamide (100 nmol/L). Only 10.8 4.2% (in HK-2 cells) or 7 1% (in PTECs) of flutamide + T treated cells showed apoptotic features at 48 hours (P < 0.01 to 0.001 vs. T, and P = NS vs. C) (Figure 5a and b).

Figure 5.

Apoptosis detection by annexin V/propidium iodide. Subconfluent proximal human tubule cell line HK-2 (A) and proximal tubular epithelial cells (PTECs) (B) were growth factor– and serum-deprived for 48 hours in the presence of testosterone (T) (10 nmol/L), flutamide (F) (100 nmol/L) alone, or T plus F (added 4 hours earlier than T). Data are shown as the mean SD from three representative experiments, with two wells of the cells for each treatment. *P < 0.01 vs. 10 nmol/L T; P < 0.001 vs. C; **P < 0.001 vs. 10 nmol/L T.

Full figure and legend (12K)

To examine whether these effects were specific to androgens, in a separate setting of experiments, cells were treated with 17-estradiol (10 nmol). Estradiol, per se, had no effects on apoptosis, but the coincubation of T + estradiol prevented the increase in the apoptosis induced by T both in HK-2 cells Figure 6a and PTECs (Figure 6b – e).

Figure 6.

Apoptosis detection by annexin V/propidium iodide. Subconfluent proximal human tubule cell line (HK-2) (A) and proximal tubular epithelial cells (PTECs) (B) were growth factor– and serum-deprived for 48 hours in the presence of T (10 nmol/L), 17-estradiol (E) (10 nmol/L) alone, or T plus E. Data are shown as the mean SD from three representative experiments, with two wells of the cells for each treatment. *P < 0.001 vs. control (C); ^#P < 0.001 vs. E; P < 0.01 vs. E. Apoptosis in primary PTEC cultures stained with annexin V (green fluorescence) and propidium iodide (red fluorescence): Control PTECs (C); PTECs treated with 10 nmol/L for 48 hours (D); PTECs exposed to T and E (10 nmol/L) for 48 hours (400) (E).

Full figure and legend (25K)

Effect of T on Fas, FasL, and FADD proteins

As a following step, we studied the expression of possible apoptosis-inducing proteins in HK-2 cells in order to individuate the underlying apoptotic pathway(s). Because of the evidence of the Fas-FasL system in proximal tubule cells¹, we first looked at the expression of Fas, FasL, and FADD induced by T treatment. Fas, a 45-kD member of the tumor necrosis factor (TNF) receptor superfamily, binds to its cognate ligand, FasL. Ligation of Fas causes rapid death-inducing FADD signaling complex formation. The protein is recruited to the receptor's DD and primes the apoptotic pathway¹. HK-2 cells were analyzed by immunofluorescence, SDS-PAGE, and Western blotting with antibodies to Fas, FasL (a 37-kD protein), and FADD protein (24-kD). The basal low amount of FasL in the cell surface is consistent with data from murine tubular cells¹⁵, as well as other cell types where most FasL is bound to intracellular membranes²². Our study revealed a clear up-regulation of Fas, FasL, and FADD in proximal tubule cells within 48 hours of T treatment Figure 7 and Figure 8

Figure 7.

Testosterone (T) treatment induces Fas (D), FasL (E), and Fas-associating death domain containing protein (FADD) (F), with respect to untreated cells (A, B, and C for Fas, FasL, and FADD, respectively) (400).

Full figure (99K)

Figure 8.

Testosterone (T)-induced apoptosis is associated with up-regulation of Fas, Fas-L, and Fas-associating death domain containing protein (FADD). Cells incubated for 48 hours with 10 nmol/L of T were subjected to Western analysis with the various antibodies as shown. The results are representative of 3 different experiments.

Full figure and legend (18K)

Effect of caspase inhibition

Starting from the above data, we sought to evaluate the downstream effectors of the Fas-FasL pathway, including the caspase family²³. Two main pathways leading to apoptosis through caspase activation have been described. On one way, the "extrinsic pathway" is activated by receptors of the TNFR superfamily through mechanisms that involve the adapter protein FADD, which activates the initiator caspase-8. On the other way, the "intrinsic pathway" is activated by mitochondial proteins through the action of the initiator caspase-9. Since the inhibition of caspase functions has been shown to block the development of programmed cell death²⁴, we evaluated whether T-induced HK-2 cell apoptosis could be prevented by treatment with different inhibitors. The results revealed that the change in cellular morphology induced by T at 48 hours were no longer observed after the addition of Z-IEDT-FMK (caspase-8 inhibitor II), Z-LEHD-FMK (caspase-9 inhibitor I) and DEVD-CHO (caspase-3 inhibitor I) to the culture medium. The antiapoptotic effects were of similar entity for all inhibitors Figure 9.

Figure 9.

Effects of the caspase inhibitor 3 (DEVD-CHO), caspase inhibitor 8 (Z-IEDT-FMK), and caspase inhibitor 9 (Z-LEHD-FMK) (50 mol/L) on 10 nmol/L testosterone (T)-induced apoptosis at 48 hours. Apoptotic cells were assessed by annexin V/propidium iodide and the values are the mean SD of 3 different experiments. *P < 0.01 vs. 10 nmol/L T.

Full figure and legend (8K)

T-induced apoptosis is associated with the activation of caspase-3

Caspase-3 is one of the key executioners of apoptosis and is responsible either partially or totally for the proteolytic cleavage of many key proteins, such as the nuclear enzyme PARP-1. HK-2 cells incubated for 48 hours with 10 nmol/L T were analyzed by immunofluorescence, SDS-PAGE, and Western blot with a polyclonal antibody to caspase-3, which detects only the p17 fragment of active caspase-3. Results showed that caspase-3 is activated during T treatment Figure 10a and b,11. We next checked for the cleavage of PARP-1, an endogenous substrate for caspase-3. During apoptosis, caspase 3 cleaves PARP-1 to yield an 85-kD and a 25-kD fragment. We examined PARP-1 expression by immunostaining and Western blot Figure 11. Cleavage into an 85-kD terminal breakdown product was detectable on Western blots in T-treated cells, confirming that caspase-3 was acting on cellular substrates.

Figure 10.

Expression of p17 fragment active caspase-3 by immunofluorescence (upper panel) and cleaved poly(ADP-ribose) polymerase (PARP-1) by immunocytochemistry (lower panel) in control (A, C) and testosterone (T)-treated cells (B, D). T activates caspase-3 and its activation is documented by PARP-1 cleavage (400).

Full figure and legend (35K)

Figure 11.

Testosterone (T)-induced apoptosis is associated with activation of caspase-3 and cleavage of poly(ADP-ribose) polymerase (PARP-1). Cells incubated for 48 hours with 10 nmol/L of T were subjected to Western analysis with the various antibodies as shown. The caspase-3 antibody detects only the p17 fragment active; the PARP antibody detects only the cleaved 85-kD fragment. The results are representative of 3 different experiments.

Full figure and legend (13K)

Effects of T on Bax and Bcl-2

Because T-induced apoptosis is inhibited by caspase-9 inhibitor I (Z-IEDT-FMK), we examined the expression of Bax and Bcl-2. T treatment significantly increased the expression of Bax protein (P < 0.01 vs. C) (Figure 12b and d). The pretreatment with flutamide did not induce this increase, and the expression of Bax was similar to control. The expression of Bcl-2 was opposite respect to Bax (Figure 12a and c).

Figure 12.

Expression of antiapoptotic Bcl-2 (A) and proapoptotic Bax (B) proteins were evaluated by immunocytochemistry and image analysis. Cells subjected to 10 nmol/L testosterone (T) for 48 hours showed alteration in Bcl-2 and Bax proteins with respect to control (C) cells and cells pretreated with flutamide (F). Data are expressed as mean SD of 4 separate experiments. *P < 0.001 vs. C. Expression of antiapoptotic Bcl-2 (C) and proapoptotic Bax (D) proteins evaluated by Western blot. Results are representative of 3 different experiments.

Full figure and legend (26K)

DISCUSSION

The present study shows that androgens promote apoptotic damage in human proximal tubule cells, as evaluated by multiple apoptosis-associated determinations, including morphologic observation of TUNEL-stained cells, annexin V binding, and detection of apoptosis-related proteins. Moreover, these effects are observed at T levels that are in the physiologic range and are blocked by androgen receptor inhibition and estrogens. These findings are observed both in immortalized and primary proximal tubular cells. According to our data, androgens increase the permissiveness of tubular cells to apoptotic effects by triggering apoptotic pathways involving Fas up-regulation, FasL expression, and caspase(s) activation.

When we blocked survival signals with cycloheximide, apoptosis occurred rapidly. We observed that combined treatment with androgens and cycloheximide generated additive apoptotic effects compared to T alone, suggesting a correlation between the inhibition of protein synthesis and T-mediated apoptosis. Both conditions induced the typical morphologic findings and the cell surface program characteristic for apoptosis.

Fas has been recognized in several nonlymphoid organs, including the kidney^1,14,24. In contrast, FasL, a 40-kD type II membrane protein, is expressed in a more restricted number of tissues, where it can promote apoptosis through the activation of Fas receptors¹. The regulation of FasL-induced apoptosis is complex, and some cell types express FasL but are unable to promote apoptosis²⁵. Fas is expressed in renal tubular epithelial cells, but the role of the Fas-FasL system on apoptosis has been demonstrated only recently¹. It has been previously observed that Fas/FasL-induced apoptosis promotes parenchymal cell damage in models of acute renal failure and glomerular injury¹. According to our data, human proximal tubular cells are activated by T and show an up-regulated expression of Fas/FasL, which can contribute to their increased susceptibility to FasL-mediated apoptosis. In keeping with these findings, we found that the effector caspase-3 was activated by T, and that a hallmark caspase substrate, PARP-1, was cleaved. As an additional finding, our data suggest that T works primarily through a FADD- and caspase-8–dependent pathway. However, the involvement of caspase-9 in the degeneration process was somewhat unexpected. On one hand, it is possible that caspase inhibitors studied here are not equipotent, but that their effect may change according to individual substrate and inhibition kinetics. On the other hand, apoptotic changes were associated with an increased expression of Bax together with a decrease in the antiapoptotic protein Bcl-2 in T-treated cells, therefore suggesting an interaction between the extrinsic and intrinsic pathway. Caspase-8 may either directly activate procaspase-3 or cleave the proapoptotic protein Bid, which then subsequently induces cytochrome c release²⁷. Nevertheless, the final result of either pathway is caspase activation, resulting in the biochemical and morphologic changes associated with the apoptotic phenotype.

The mechanisms responsible for gender differences in progression rates of chronic diseases are not yet clear and continue to be the object of investigation. Changes observed in tubule cells under serum-free culture in vitro appear very similar to the apoptotic damage occurring in vivo in the tubuli subjected to ischemia or chronic renal disease^28,29,30. The androgen-promoted apoptotic damage might interact with the apoptotic pathway already activated in several chronic kidney diseases. It has been previously shown that renal FasL may potentially regulate the immune response and promote parenchymal cell death in several experimental models of chronic kidney disease²². The Bax/Bcl-2 ratio (through caspase-3 activation) has been shown to modulate renal cell apoptosis associated with the progressive renal damage in immune-mediated experimental glomerulonephritis²⁶, in the subtotal nephrectomy model of chronic renal scarring²⁹, in adult polycystic kidney disease (APKD)³⁰, and in experimental obstructive nephropathy³¹. Therefore, the pathways to cell death, which are primed by androgens, as shown by the present study, are shared by several conditions leading to the loss of renal cells in chronic disease.

CONCLUSION

Physiologic T levels cause intracellular events, allowing Fas sensitization and leading to activation of caspase-dependent apoptotic pathway in human renal tubule cells. This is consistent with a role for T in promoting renal injury in men.

References

References

1.	ORTIZ A. Nephrology forum: Apoptotic regulatory proteins in renal injury. Kidney Int 2000; 58 1: 467–485. | PubMed |
2.	GILL C, MESTRIL R & SAMALI A. Losing heart: The role of apoptosis in heart disease—A novel therapeutic target?. FASEB J 2002; 16 2: 135–146. | Article | PubMed | ChemPort |
3.	PADANILAM BJ. Cell death induced by acute renal injury: A perspective on the contributions of apoptosis and necrosis. Am J Physiol Renal Physiol 2003; 284 4: F608–627. | PubMed |
4.	GENNARI A, CORTESE E & BOVERI M et al. Sensitive endpoints for evaluating cadmium-induced acute toxicity in LLC-PK1 cells. Toxicology 2003; 183 1–3: 211–220. | PubMed |
5.	SILBIGER RS & NEUGARTEN J. The impact of gender on the progression of chronic renal disease. Am J Kidney Dis 1995; 25 4: 515–533. | PubMed |
6.	ANTUS B, HAMAR P & KOKENY G et al. Estradiol is nephroprotective in the rat remnant kidney. Nephrol Dial Transplant 2003; 18 1: 54–61. | PubMed |
7.	NEUGARTEN J. Gender and the progression of renal disease. J Am Soc Nephrol 2002; 13: 2807–2809. | PubMed |
8.	NEGULESCU O, BOGNAR I & LEI J et al. Estradiol reverses TGF-1–induced mesangial cell apoptosis by a protein kinase CK2-dependent mechanism. Kidney Int 2002; 62 6: 1989–1998. | Article | PubMed |
9.	BALTATU O, CAYLA C & ILIESCU R et al. Abolition of hypertension-induced end-organ damage by androgen receptor blockade in transgenic rats harboring the mouse ren-2 gene. J Am Soc Nephrol 2002; 13 11: 2681–2687. | PubMed |
10.	FISCHER GM, CHERIAN K & SWAIN ML. Increased synthesis of aortic collagen and elastin in experimental atherosclerosis: Inhibition by contraceptive steroids. Atherosclerosis 1981; 39: 463–467. | PubMed | ChemPort |
11.	LING S, DAI A & WILLIAMS MR et al. Testosterone enhances apoptosis-related damage in human vascular endothelial cells. Endocrinology 2002; 143 3: 1119–1125. | PubMed |
12.	TESARIK J, MARTINEZ F & RIENZI L et al. In-vitro effects of FSH and T withdrawal on caspase activation and DNA fragmentation in different cell types of human seminiferous epithelium. Human Reprod 2002; 17: 1811–1819.
13.	MIYAKE H, NELSON C & RENNIE PS et al. Testosterone-repressed prostate message-2 is an anti-apoptotic gene involved in progression to androgen independence in prostate cancer. Cancer Res 2000; 60 1: 170–176. | PubMed | ISI | ChemPort |
14.	BOONSTRA JG, VAN DER WOUDE FJ & WEVER PC et al. Expression and function of Fas (CD95) on human renal tubular epithelial cells. J Am Soc Nephrol 1997; 8: 1517–1524. | PubMed | ISI | ChemPort |
15.	LORZ C, ORTIZ A & JUSTO P et al. Pro-apoptotic Fas ligand is expressed by normal kidney tubular epithelium and injured glomeruli. J Am Soc Nephrol 2000; 11 7: 1266–1277. | PubMed | ISI | ChemPort |
16.	THOMAS GL, YANG B & WAGNER BE et al. Cellular apoptosis and proliferation in experimental renal fibrosis. Nephrol Dial Transplant 1998; 13: 2216–2226. | Article | PubMed | ISI | ChemPort |
17.	SUGIYAMA H, KASHIHARA N & MAKINO H et al. Apoptosis in glomerular sclerosis. Kidney Int 1996; 49 1: 103–111. | PubMed | ISI | ChemPort |
18.	DETRISAC CJ, SENS MS & GARVIN AJ et al. Tissue culture of human kidney epithelial cells of proximal tubule origin. Kidney Int 1984; 25: 383–390. | PubMed | ISI | ChemPort |
19.	VERZOLA D, BERTOLOTTO MB & VILLAGGIO B et al. Taurine prevents apoptosis induced by high ambient glucose in human tubule renal cells. J Investig Med 2002; 50 6: 443–451. | PubMed | ISI | ChemPort |
20.	VERZOLA D, VILLAGGIO B & BERRUTI V et al. Apoptosis induced by serum withdrawal in human mesangial cells: Role of IGFBP-3. Exp Nephrol 2001; 9 6: 366–371. | PubMed |
21.	LAPOSATA M. The New England Journal of Medicine SI Unit Conversion Guide 1992; Boston, NEJM Books.
22.	ORTIZ A, LORZ C & JUSTO P et al. Contribution of apoptotic cell death to renal injury. J Cell Mol Med 2001; 5 1: 18–32. | PubMed |
23.	SMITH CA, FARRAH T & GOODWIN RG. The TNF receptor superfamily of cellular and viral proteins: Activation, costimulation, and cell death. Cell 1994; 76: 959–962. | Article | PubMed | ISI | ChemPort |
24.	NICHOLSON DW. ICE/CED-3-like proteases as therapeutic targets for the control of inappropriate apoptosis. Nat Biotech 1996; 14: 297–301. | PubMed |
25.	ITOH N, YONEHARA S & ISHLL A et al. The polypeptide encoded by the cDNA for human cell surface antigen Fas can mediate apoptosis. Cell 1991; 66: 233–243. | Article | PubMed | ISI | ChemPort |
26.	NAGATA S. Apoptosis by death factors. Cell 1997; 88: 355–36. | Article | PubMed | ISI | ChemPort |
27.	LUO X, BUDIHARDJO I & ZOU H et al. Bid, a Bcl-2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors. Cell 1998; 94: 481–490. | Article | PubMed | ISI | ChemPort |
28.	YANG B, JOHNSON TS & THOMAS GL et al. A shift in the Bax/Bcl-2 balance may activate caspase-3 and modulate apoptosis in experimental glomerulonephritis. Kidney Int 2002; 62 4: 1301–1310. | Article | PubMed |
29.	YANG B, JOHNSON TS & THOMAS GL et al. Expression of apoptosis-related genes and proteins in experimental chronic renal scarring. J Am Soc Nephrol 2001; 12 2: 275–288. | PubMed | ISI | ChemPort |
30.	WOO D. Apoptosis and loss of renal tissue in polycistic kidney disease. N Engl J Med 1995; 333: 18–25. | Article | PubMed | ISI | ChemPort |
31.	ZHANG G, OLDROYD SD & HUANG LH et al. Role of apoptosis and Bcl-2/Bax in the development of tubulointerstitial fibrosis during experimental obstructive nephropathy. Exp Nephrol 2001; 9 2: 71–80. | Article | PubMed | ISI | ChemPort |

Acknowledgments

This study was supported by grants from the University of Genoa (Cofinanziamento di Ateneo) and the Italian Ministero dell'Università e della Ricerca Scientifica (FIRB project). The cost of color pages was supported by Dompé Biotech, Milan, Italy.