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Original Studies |
Istituto di Endocrinologia (Da.P., V.R., A.B., A.A.S.), Dipartimento di Biochimica e Biofisica (V.G.), Seconda Universitá di Napoli; Istituto di Urologia, Universitá Federico II (D.P., T.L.); and Dipartimento di Biochimica e Biotecnologie Mediche-Centro di Ingegneria Genetica, Università Federico II (V.C.), 80131 Naples, Italy
Address all correspondence and requests for reprints to: Antonio A. Sinisi, M.D., Istituto di Endocrinologia, Seconda Universitá di Napoli, Building 16, Via Pansini 5, 80131 Naples, Italy. E-mail: sinisi{at}unina.it
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Abstract |
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, -ß, and -
), tTGase, retinol-binding protein
(RBP), and cellular RBP type I transcripts were determined by
semiquantitative RT-PCR. After 72–96 h of 10-6 mol/L RA
treatment, cell growth inhibition and apoptosis were associated with
increased tTGase activity in both NEPC and CEPC, and with increased
tTGase protein and messenger ribonucleic acid levels only in NEPC.
Moreover, RA down-regulated RAR and -ß and increased RBP messenger
ribonucleic acid levels in NEPC, whereas it increased RARß gene
expression and decreased Bcl-2 protein levels in CEPC. Our results
suggest that RA induces tTGase gene expression and enzyme activity in
normal prostate cells, and that RA-regulated pathways are impaired in
cancer cells. Moreover, down-regulation of Bcl-2 protein and
up-regulation of RARß suggest that retinoid may act on the genetic
defect responsible for prostate cancer progression. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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It is not known whether prostate cancer is due to alterations of
retinoid content and/or of retinoid-regulated pathways. The effects of
retinoids are mediated by two classes of nuclear proteins, called
retinoic acid (RAR, -ß, and -) and retinoic X, -ß and -
receptors, which are ligand-regulated transcription factors (9).
Retinoids bind another family of proteins involved in their
extracellular and intracellular transport and metabolism, including the
retinol-binding protein (RBP), cellular RBP type I (CRBP-I) and type II
(CRBP-II), and two cellular RA-binding proteins (9, 10). The
antiproliferative effects of retinoids rely on the regulation of many
biological processes, including cell proliferation, differentiation,
and programmed death or apoptosis (10, 11). Hormone-induced tissue
remodeling and apoptosis are associated with changes in the products of
several genes, including tissue transglutaminase (tTGase), a
Ca2-dependent enzyme catalyzing the formation of
--glutamyl-lysine cross-links between polypeptide chains (11, 12). RA activates tTGase and modulates its expression via specific RAR
subtypes in some cell systems (12, 13), but there are no data for human
prostate cells. Furthermore, evidence suggests that the expression of
Bcl-2, an antiapoptotic protein also regulated by RA (14), is
associated with the emergence of androgen resistance and invasiveness
of prostate cancer (15, 16, 17). The aim of this study was to investigate
the effects of RA on the growth and programmed cell death of human
prostate epithelial cell (EC) primary cultures. In this context, we
evaluated whether RA regulates tTGase activity and expression and the
Bcl-2 protein level. Moreover, we studied whether the expression and
regulation of the genes mediating retinoid action, i.e.
RAR, RARß, RAR, CRBP-I, and RBP, differ between epithelial
cells from human normal prostate (NEPC) and those from
prostate cancer (CEPC).
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Normal human prostatic tissues were collected from patients who had undergone radical cystectomy for bladder cancer. Prostate cancer tissues were obtained from patients who had undergone radical prostatectomy (Gleasons’ score, 6–8). After prostatectomy, a wedge-shaped specimen of the fresh prostate was removed. A sample of the tissue underwent pathological examination to confirm prostatic origin, diagnosis, and absence of other diseases. Only specimens containing 100% normal or cancer prostate cells were used to establish primary cultures according to a previously described method (18). Prostate EC were separated by different centrifugations of minced and collagenase (collagenase IV, Life Technologies, Milan, Italy; 10 mg/mL)-digested tissues. The EC were plated on serum-free keratinocyte medium (Life Technologies) supplemented with bovine pituitary extract (10 mg/mL), epidermal growth factor (10 ng/mL), cholera toxin (10 ng/mL), 5% FCS, and antibiotics (fungizone and penicillin-streptomycin). At confluence, cultures were grown after ethylenediamine tetraacetate (EDTA)-trypsin treatment. At the first passage, immunocytochemical staining with monoclonal antibodies specific for cytokeratin (anticytokeratin Pan, Boehringer Mannheim, Mannheim, Germany) showed that the EC cultures were nearly 100% pure. Both normal and malignant primary cell cultures showed positive immunoreactivity with cytokeratin 8 monoclonal antibody (35BH11, Dako Corp., Milan, Italy), indicating their glandular origin. High mol wt cytokeratin immunoreactivity (MAb 35BE12, Dako Corp.) was completely absent from the cells derived from prostate carcinoma, but was present in cultures of normal cells, demonstrating the presence of cells endowed with basal features. The detection of prostate-specific antigen protein in conditioned medium by specific immunoassay and the immunoreactivity of cell monolayers to MAb clone ErPr8 (BioGenex, San Ramon, CA) demonstrated that both normal and malignant short term primary cell cultures retain prostate-specific antigen secretory function. The malignant nature of cells derived from prostate carcinomas was confirmed by a high expression of the proliferative antigen Ki-67 and particularly by a high expression of mutated p53 protein, demonstrated by immunoreactivity with monoclonal antibodies clone Ki-67 (Dako Corp.) and PAb 240 (Serotec, Delta Biological, Italy), respectively. Four cell strains from normal prostates and four from prostate cancer specimens were used in the experimental protocols, which were repeated at least three times. All cultures were performed at 37 C in a humidified 5% CO2 atmosphere.
Morphological evaluation
The cells from primary monolayers were detached with trypsin-EDTA and seeded in 60-mm culture dishes to grow at 70–80% confluence. Cells were starved for 24 h in MEM without FCS and incubated in 1% FCS-supplemented medium in the presence of RA (10-6, 10-7, and 10-8 mol/L) or solvent (control cells) for 96 h. Morphology was assessed with an inverted microscope (Nikon Duofot, Nikon, Tokyo, Japan).
Cell proliferation assay
Cell proliferation was evaluated with the MTT method (Boehringer Mannheim). Cells were seeded in microtiter plates in a final volume of 100 µL complete culture medium at a concentration of 2 x 103 cells/well and grown for 24 h at 37 C in 5% CO2. Cells, starved for 24 h in MEM without FCS, were incubated in 1% FCS-supplemented medium in the presence of 10 -6, 10-7, and 10-8 mol/L RA or solvent (control cells) for 96 h. Then, 10 µL MTT solution were added to each well, and plates were further incubated for 4 h. Ten microliters of solubilization solution were added to each well, and plates were kept overnight in the incubator. The absorbency was read at 550 nm using a microtiter plate reader.
Apoptosis detection
The In Situ Cell Death Detection Kit (Boehringer Mannheim; TUNEL) was used to detect apoptosis and quantify DNA strand breaks in individual cells. The cell monolayers were grown directly on sterilized slides (SuperFrost, Carlo Erba, Milan, Italy), starved for 24 h in MEM without FCS, and then incubated in 1% FCS-supplemented medium in the presence of RA (10-6, 10-7, and 10-8 mol/L) or solvent (control cells) for 96 h. The slides were then fixed in buffered paraformaldehyde permeabilized with Triton-X and labeled with TUNEL reaction mixture according to the manufacturer’s instructions. Samples were analyzed using a Leitz Diaplan microscope (Leica, Milan, Italy) equipped with epifluorescence. A negative control (obtained by incubating a slide with labeled solution without terminal transferase) and a positive control (obtained by treating a slide with deoxyribonuclease I solution) were included in each assay run.
tTGase assay
The cells were washed three times with PBS, scraped with a rubber policeman from the dishes in the presence of TE (1 mmol/L EDTA in 10 mmol/L Tris-HCl, pH 7.4), centrifuged, resuspended at a final concentration of 5 x 106 in the same buffer, and then sonicated at 4 C for 20 s. The tTGase activity of the sonicate was assessed by measuring the [14C]spermidine ([14C]Spd) incorporation into N,N'-dimethylcasein. The incubation mixture contained, in a final volume of 100 µL, 100 mmol/L Tris- HCl (pH 7.5), 10 mmol/L dithiothreitol, 0.25 µCi [14C]Spd (Amersham, Milan, Italy; 101 mCi/mmol), 40–80 µg sonicate protein, and, where indicated, 2.5 mmol/L CaCl2, 20 mg/mL N,N'-dimethylcasein, and 5 mmol/L ethyleneglycol-bis-(ß-aminoethyl ether)-N,N,N',N'-tetraacetic acid. After 30 min of incubation at 37 C, the reaction was stopped by the addition of 10 µL 1 mol/L unlabeled Spd and 1 mL ice-cold 10% trichloroacetic acid (TCA). The TCA-precipitated material, washed three times with 1 mL cold 10% TCA, was dissolved in 100 µL 1 mol/L NaOH, and its radioactivity was measured in a Packard liquid scintillation counter (Downers Grove, IL).
Electrophoresis and Western blot (WB) analysis
tTGase, P53, p21, and Bcl-2 protein levels were evaluated by WB analysis of protein extracts made from three different strains of NEPC and CEPC. For electrophoresis and WB analysis, the cells were harvested after a few minutes of incubation with PBS containing 0.2 mmol/L EDTA and centrifuged, and the pellets containing 10-6 cells were resuspended in 2 x denaturing lysis buffer (1:1, vol/vol) containing 0.25 mol/L Tris-HCl (pH 6.8), 5% SDS, 8 mol/L urea, 10 mmol/L EDTA, and 0.1 mol/L dithiothreitol. The cell lysates were centrifuged for 10 min at maximum speed at room temperature to separate DNA, and then the supernatant was boiled for a few minutes before loading on gels. Protein concentrations were normalized, and equal volumes of samples were loaded on the gels. Electrophoresis was performed on 12% polyacrylamide (1:40, mono/bis acrylamide) containing SDS according to usual SDS-PAGE procedures. After separation on gel, proteins were electrophoretically transferred overnight to 0.45-µm nitrocellulose sheets for WB analysis in transferring buffer containing 20% methanol, 10 g/L glycine, 4 g/L Tris, and 0.2 g/L SDS. Nitrocellulose reactive groups were then blocked with WB buffer (3 g/L Na2HPO4, 0.3 g/L NaH2PO4, 12 g/L NaCl, 0.05% Nonidet P-40, and 0.05% Tween-20) containing 4% nonfat dried milk (Blocker, Bio-Rad, Rome, Italy) and 1% BSA (pH 8.00). After 1 h of incubation with blocking solution at room temperature, the sheets were briefly washed with WB buffer (pH 8.00), incubated overnight, and shacked at 4 C with primary antibodies diluted in WB buffer containing 1% nonfat dried milk and 0.25% BSA (pH 8.00). We used the following antibodies for the primary immunoreactions: a goat affinity-purified polyclonal antibody to tTGase at a working concentration of 1:2000, for p53 we used the p53 PAN 122 (Boehringer Mannheim, Florence, Italy) at a working concentration of 2 µg/mL, for Bcl-2 we used the monoclonal antibody (Boehringer Mannheim) at a working concentration of 1:800, and for p21WAF we used the monoclonal antibody Ab1 (Oncogene Research Science, Cambridge, MA) at a working concentration of 0.5 µg/mL. At the end of incubation, blots were washed once for 15 min and three times for 5 min each time with WB buffer (pH 8.00). Antibody reaction was revealed by incubation for 45 min at room temperature with horseradish peroxidase-coupled antigoat or antimouse IgG serum (Amersham, Milan, Italy), 1:10,000 diluted in WB buffer (pH 8.00) containing 1% nonfat dried milk and 0.25% BSA, followed by a washing cycle (as described above) and using chemiluminescent substrate (ECL, Amersham) according to the manufacturer’s instructions. The visualization was obtained by autoradiography.
Ribonucleic acid (RNA) isolation
RNA was isolated from the cultures at the first passage (for EC). Total RNA was recovered with the RNAzol B kit (Cinna/Biotecx Laboratories, Houston, TX). Residual DNA was removed by ribonuclease-free deoxyribonuclease I treatment (Promega Corp., Florence, Italy).
RT-PCR
RNAs were reversely transcribed using 5 µg total RNA as
previously described (5, 19). To obtain a negative control for the
amplification reactions, we carried out RNA transcription without the
addition of reverse transcriptase. Four hundred nanograms of
complementary DNA obtained by RT of RNAs were amplified in the total
volume of 50 µL 10 mmol Tris-HCl, 1.5 mmol MgCl2, 50 mmol
KCl (pH 8.3), and 100 ng 5'-3'-end primers. PCR conditions were as
previously described, and the reaction consisted of 32 cycles of
amplification (19). To determine whether the expression of RARs and
CRBP-I, RBP, and tTGase varied between individual tissue samples, a
semiquantitative PCR was performed in which these genes were amplified
with glyceraldehyde-3-phosphate dehydrogenase (GAPDH; 22 cycles) as
previously described (19). The number of cycles was chosen in the
middle of the exponential phase of the reaction, separately for each
type. To establish the number of cycles, GAPDH was amplified at 15, 22,
32, and 40 PCR cycles; RAR, -ß, and - were subjected to 25, 32,
and 40 amplification cycles (data not shown). The levels of messenger
RNAs (mRNAs), quantified by densitometric scanning of the amplification
products electrophoresed on agarose gels, are expressed as the ratio
between the density of each gene product and that of coamplified GAPDH.
We used oligonucleotide sequences for RAR, RARß, RAR, CRPB-I,
and GAPDH as previously described (5, 19). We also used the following
primers: tTGase sense, 5'-GCATGGTCAACTGCAACGATG-3'; and antisense,
5'-GGGCGCATCGTACTTGGTG-3'; and RBP sense,
5'-GCCAAGAAGGACCCCGAGGGCCTC-3'; and antisense,
5'-ATTTCCTTTCTGCAGAAAGGAGGC-3'. PCR products were then separated on a
1.2% agarose gel containing ethidium bromide using a 100-bp DNA ladder
(Life Technologies, Milan, Italy) as a size marker.
Statistical analysis
The data are reported as the mean ± SEM obtained from at least three separate experiments in which each point was performed in triplicate. The means were compared using ANOVA.
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Results |
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NEPC and CEPC were incubated for 96 h with RA at
10-6, 10-7, and 10-8 mol/L.
Morphological changes were observed after 72 and 96 h of
10-6 mol/L RA treatment in NEPC and CEPC, respectively.
Cells became thinner and oblong, and showed cytoplasm extensions
projecting from their surface; the cytoplasm contained numerous
vacuoles (Fig. 1
). Treatment with
10-6 mol/L RA inhibited cell proliferation and viability,
as measured by the MTT method; inhibition was significant after 72
h in both NEPC and CEPC (P < 0.0001 and
P < 0.02, respectively, vs. untreated
controls; Fig. 2). DNA nuclear
fragmentation was detected (TUNEL method) in 10% of NEPC cells and in
2% of CEPC cells treated with 10-6 mol/L RA for 96
h, but not in untreated control cultures (Fig. 3).
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As shown in Table 1, tTGase activity
was induced in a dose-dependent manner by RA treatment in NEPC and
CEPC; tTGase activity was maximal at a dose of 10-6 mol/L
after 72 h (P < 0.0001 and < 0.005,
respectively).
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The tTGase protein level, determined by WB in cell lysates,
increased in NEPC treated with 10-6 mol/L RA for 72 h
(Fig. 4). To test whether the p53 pathway
was functionally activated during RA treatment, we determined whether
p53 and p21 proteins were present in EC and whether their levels
increased after RA administration. In untreated NEPC, a ladder of low
mol wt protein bands was recognized by the monoclonal p53 antibody,
showing that wild-type p53 is degraded; after treatment with RA for
72 h, the ladder of bands was unrecognized, and there was a
selective increase in the labeling of the band corresponding to the
wild-type p53 (Fig. 5). The p21 protein
level was very low in NEPC both before and after 72-h RA treatment
(data not shown). RA did not affect p53 and p21 levels in CEPC (Fig. 6). Cell extracts were also analyzed using a monoclonal Bcl-2 antibody.
In NEPC, Bcl-2 expression was undetectable both before and after RA
treatment (data not shown); in CEPC, the Bcl-2 protein level, detected
under basal conditions, was negligible after RA treatment (Fig. 6).
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, RARß, RAR, RBP, CRBP-I, and tTGase are expressed in
NEPC and CEPC, and RAR, RARß, tTGase, and RBP mRNA levels are
modulated by RA
To test whether RA induced the expression of RAR, RARß,
RAR, CRBP-I, RBP, and tTGase genes in our in vitro
system, we used a semiquantitative RT-PCR procedure to evaluate the
level of their transcripts after administration of 10-6
mol/L RA, i.e. the dose most effective in inducing
morphological and biochemical changes. Figures 7 and 8
show the transcript levels in NEPC and CEPC at 72 h after RA
administration. The basal expression of all of the genes tested was
comparable in normal and malignant cells. After 72 h of treatment,
RAR and RARß mRNA levels decreased (P < 0.007),
and tTGase and RBP mRNA levels increased (P < 0.01) in
NEPC, whereas the RARß level increased in CEPC (P <
0.001).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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- glutamyl-lysine cross-links between
polypeptide chains in the apoptotic bodies (11, 23, 24). In an attempt to clarify the mechanisms involved in RA-induced apoptosis in prostate cells, we also examined the expression of other apoptosis-related proteins, i.e. p53, p21, and Bcl-2. RA induced an accumulation of p53 in NEPC, inhibiting the appearance of the degraded forms of the protein; however, the appearance of apoptosis in our cells does not seem to involve a p53-regulated G1 arrest, because p21 protein levels remained unchanged, as observed also in nonproliferating normal prostate cells deprived of androgen (25). Immunodetection in CEPC of Bcl-2 protein, which negatively influences the apoptotic event, demonstrates that these cell types derive from advanced and hormone refractory prostate adenocarcinomas (15, 16, 17, 26). Down-regulation of Bcl-2 levels in CEPC suggests that RA exerts a direct regulation of Bcl-2 protein in prostate cancer cells, as occurs in human leukemic cells (14).
The effects exerted by RA on cancer cell growth control and apoptosis
are mediated by specific RAR subtypes and are modulated by the tissue
levels of binding proteins (1, 9, 10). The increase in tTGase
associated with retinoid-regulated apoptosis appears to be specifically
induced via the RAR subtype in rat tracheo-bronchial cell lines
(12). RARs may play a role in controlling cell proliferation and
apoptosis, and the aberrant expression of one or more RARs could result
in abrogated retinoid signaling and increased cell transformation in
several cancer types, including prostate, breast, and lung cancers (21, 22, 27).
We show that RAR and -ß mediate retinoid action in primary
prostate cell cultures, with some differences between the two cell
types examined at 72 h of RA treatment: RAR and -ß mRNAs were
down-regulated in NEPC, whereas RARß mRNA was increased. This finding
and our unpublished observation of increased RARß mRNA 5–24 h
post-RA in NEPC point to abnormalities in the regulation of RARß
expression and function in prostate cancer cells, which are partially
counteracted by RA. Experimental data show that retinoids modulate
their own metabolism by regulating the expression of RBP and CRBP genes
(9, 10, 28, 29). We provide the first demonstration that RBP and CRBP-I
genes are expressed in both NEPC and CEPC, a finding that confirms that
these genes are involved in RA metabolism and action at tissue and
cellular level in human prostate. Our study also demonstrates that RA
regulates the expression of RBP in human normal prostate cells, but not
in cancer cells, which is further evidence of a different retinoid
metabolic status in prostate cancer vs. normal prostate.
In conclusion, the natural retinoid RA, albeit at supraphysiological doses, exerts antiproliferative effects on human prostate normal and cancer cells, and up-regulates tTGase. Our data suggest that RA-induced tTGase activity, increased protein level, and increased gene expression are associated with the appearance of programmed cell death in normal human prostate cells. Previous and present studies suggest that scarce retinoid availability and action at the cellular level because of either decreased content or deranged metabolism in prostate cancer cells play a pivotal role in the abnormal cellular differentiation pathways, loss of antiproliferative effect, and accumulation of aberrant cells. Moreover, the down-regulation of antiapoptotic oncoprotein Bcl-2 and the up-regulation of RARß observed in CEPC support the hypothesis that retinoids may act on the deregulated mechanisms involved in the advance of prostate cancer and the appearance of hormone resistance. In the light of these in vitro data, it may be useful to evaluate whether synthetic retinoids, which are less toxic and more readily available than natural retinoids, can overcome the alterations of RA-mediated pathways in prostate cancer and, moreover, to determine whether they counteract in vivo the genetic defect responsible for hormone resistance and metastatization, thus functioning as significant chemopreventive agents.
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Acknowledgments |
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Footnotes |
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Received August 6, 1998.
Revised December 23, 1998.
Accepted December 30, 1998.
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References |
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, ß and , and cellular retinol binding
protein-I expression in breast fibrocystic disease and cancer. Eur J
Endocrinol. 137:410–414.[Abstract]
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