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Original Studies |
Institute of Reproduction and Development, Monash University (T.Z.T., H.W., P.N., M.K.O., G.P.R.), Clayton; and Melbourne Pathology (J.P.), Collingwood, Australia; and Oxford Brookes University (L.W.E., N.P.G.), Headington, United Kingdom
Address all correspondence and requests for reprints to: Dr. G. P. Risbridger, Institute of Reproduction and Development, Monash University, Level 3, Block E, Monash Medical Center, 246 Clayton Road, Clayton 3168, Australia. E-mail: gail.risbridger{at}med.monash.edu.au
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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with either ßA (inhibin A) or
ßB (inhibin B). The inhibin ßA- and
ßB-subunits can also form dimers called activins; three
forms are known to exist: activin A (ßAßA),
activin B (ßBßB), and activin AB
(ßAßB) (4). Activins have been shown to
have either proliferative or antiproliferative actions in reproductive,
neuroendocrine, and erythroid tissues (5, 6, 7). The effects of activins
on cultured cells include induction of cell cycle arrest (8) and
apoptosis (9). Follistatins are structurally unrelated proteins (which
also stimulate FSH secretion) (10) and bind activins, resulting in the
neutralization of activin bioactivity (9, 11). Follistatins are
glycosylated monomeric proteins that arise from two alternatively
spliced messenger ribonucleic acid (mRNA; FS315 and FS288), resulting
in three protein cores; nine mol wt forms are postulated, and six have
been identified and characterized (12).
Recent studies from this laboratory have shown that inhibins, activins,
and follistatins are synthesized by the human prostate gland. Thomas
and colleagues (13) have reported mRNA expression for - and
ß-subunits, the activin type II receptor, and both follistatin
transcripts in tissues from men with benign prostatic hyperplasia.
Proteins for activin A, activin ßA and ßB
subunits, and follistatins were localized to the hyperplastic human
prostate by immunohistochemical techniques (13). More recently, inhibin
-subunit gene expression and protein synthesis were reported in
benign prostate hyperplasia (BPH) tissues; however, in malignancy,
down-regulation of inhibin gene expression occurred, and no mRNA or
protein was observed in tissues from patient with high grade cancers
(Mellor SL, Richards MG, Pedersen JS, Robertson DM, and Risbridger GP,
submitted for publication). The loss of inhibin expression in
vivo correlated with the failure to detect inhibin gene or
protein in human prostate tumor cell lines (14, 15, 16). Together with the
findings that functional deletion of the inhibin -subunit results in
the development of gonadal tumors in mice (17), it was proposed that
the inhibin gene is a tumor suppressor (16).
In the absence of inhibin gene expression in high grade prostate
cancer, it remains to be determined whether there is additional
regulation of activin ß-subunits, which may involve follistatin
expression. In vitro studies using the human prostate tumor
cell lines LNCaP, DU145, and PC3 have reported activin ß-subunit and
follistatin mRNAs and proteins (14, 15, 16, 18), suggesting endogenous
production of these proteins by prostate tumor cells. However, the
effect of exogenously added activins A and B was to inhibit the growth
of LNCaP and DU145 cells; this response was not observed by PC3 cells,
which appear resistant to the effects of activin A (18). Although
activin inhibits tumor cell proliferation in vitro,
follistatin completely blocks this action on DU145 and LNCaP cells
(18). Therefore, the different in vitro observations may be
due to the production of different levels of follistatins and/or
protein forms by each of the tumor cell lines. The aim of this study
was to determine whether poorly differentiated tumor cells from men
with high grade prostate cancer express activin ßA- and
ßB-subunits and follistatins. The patterns of
localization and expression in malignant tissue were compared with
those in nonmalignant regions of the same tissue specimens.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Needle biopsy specimens were obtained from a total of 25
patients undergoing transrectal ultrasound (TRUS)-guided biopsy.
Carcinoma of the prostate was confirmed by histological examination
(Melbourne Pathology, Collingwood, Australia), and the patients
selected were those diagnosed with high grade prostate cancer and a
Gleason score between 7–10. The specimens were obtained in accordance
with the requirements and approval of the standing committee for human
ethics and experimentation at Monash Medical Center and Monash
University. Four-micron sections were cut from the specimens and used
for immunohistochemistry and/or in situ hybridization (shown
in Table 1
) as described below. The
expression or localization of inhibin ß-subunits or follistatin was
compared in nonmalignant regions and malignant (Gleason grade 4 or 5)
regions of the tissue sections.
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Antibodies. Four antibodies for the inhibin ßA- and ßB-subunit proteins were used for these studies. The ßA and ßB Salk polyclonal antibodies (gifts from Dr. W. Vale) were raised in rabbits against synthetic human ßA or ßB peptides (19). The ßA and ßB Groome monoclonal antibodies (gifts from Dr. N. P. Groome) were raised against synthetic human ßA or ßB peptides and have previously been used for the measurement of inhibin A and inhibin B (20, 21).
Follistatin immunoreactivity was determined using two antisera. AS 202 was raised in an intact adult male New Zealand rabbit to purified bovine 39-kDa follistatin and has been used for the specific measurement of follistatin in serum samples by RIA (22). On Western blot analysis, the cross-reaction of bovine follistatin with AS 202 was approximately 16 times greater than that with human recombinant FS288 per ng protein (de Kretser, D. M., unpublished observation). Preabsorption of AS 202 was achieved by incubating 1 µg bovine follistatin with 25 µL of a 1:500 dilution of AS 202 overnight at 4 C. The mixture was centrifuged at 12,000 x g, and the supernatant was collected and used accordingly. The OxB288 antibody was raised to human recombinant FS288 and was generously provided by Dr. L. Evans (23).
Immunohistochemistry. Sections were dewaxed, rehydrated, and placed in Target Retrieval Solution (Dako, Carpenteria, CA). Antigenic sites were exposed by microwaving the sections (2.25 watts/mL·min for 3 min, followed by 0.3 watts/mL·min for 3–5 min or 20 min at 2.25 watts/mL·min) before the removal of endogenous peroxidase activity with 3% H2O2 for 30 min. Sections were incubated with 0.2% Triton X-100 (Sigma Chemical Co., St. Louis MO) for 10 min before blocking in either 1% blocking reagent (Boehringer Mannheim, Mannheim, Germany) or CAS block (Zymed, San Francisco, CA) plus 10% normal serum. The sections were incubated with the primary antibody overnight at 4 C (ßA, ßB, and OxB288) or at room temperature for 2 h (AS 202), washed in PBS (10 mmol/L phosphate buffer and 15 mmol/L NaCl, pH 7.4), and incubated with either biotinylated anti-rabbit IgG (1:200; Salk antibodies and AS 202; Zymed) or biotinylated antimouse IgG (1:200; Groome antibodies; Vector Laboratories, Burlingame, CA) at room temperature for 1 h. The sections were washed in 0.01 mol/L PBS and incubated for 1 h with streptavidin (1:50; Vectastain Elite ABC Kit, Vector Laboratories). After additional washes with 0.01 mol/L PBS, peroxidase activity was detected using 3',3'-diaminobenzidine tetrahydrochlorate (Liquid DAB Substrate Kit, Zymed). The reaction was terminated in distilled water, and the sections were counterstained with Mayer’s hematoxylin (Sigma Diagnostics, St. Louis, MO), dehydrated, and permanently mounted with DPX (BDH, Poole, U.K.).
In situ hybridization studies
Probe synthesis. Digoxygenin (DIG)-labeled riboprobes were prepared using the Boehringer Mannheim riboprobe labeling kit. Sense and antisense complementary RNA (cRNA) probes were generated from rat inhibin ßA and ßB (24) and rat follistatin (25) partial clones (370, 390, and 267 bp, respectively).
Hybridization. Sections were dewaxed, rehydrated, treated with 0.2 mol/L HCl, and then washed in diethyl pyrocarbonate-treated water (twice, 5 min each time). Sections were digested with proteinase K (Boehringer Mannheim) for 30 min at 37 C. After incubation in 0.2% glycine for 10 min at 4 C, sections were equilibrated in 0.1 mol/L triethanolamine and acetylated in 0.25% acetic anhydride for 5 min. After rinsing in diethyl pyrocarbonate-treated water, sections were prehybridized for a minimum of 30 min. Prehybridization solution contains 3 x SSC (1 x SSC is 0.15 mol/L sodium chloride and 15 mmol/L sodium citrate, pH 7), 1 x Denhardt’s solution, 50% deionized formamide, 66 mmol/L phosphate buffer (pH 8), 1000 µg/mL herring sperm DNA, and 200 µg/mL transfer RNA.
Sections were hybridized overnight at 42 C in hybridization buffer (prehybridization solution plus 10% dextran sulfate) containing a predetermined concentration (200–1000 ng/mL) of riboprobe. Excess probe was removed by sequential 15-min washes in 2 x SSC at room temperature, 2 x SSC at 42 C, 1 x SSC at 42 C, and finally either 0.1 or 0.5 x SSC (depending on the probe employed) at 42 C.
Anti-DIG detection. The tissues were briefly washed in buffer 1 (0.1 mol/L maleic acid and 0.15 mol/L sodium chloride, pH 7.5) before anti-DIG antibody detection. Anti-DIG Fab fragments conjugated to alkaline phosphatase (Boehringer Mannheim) were diluted to 1:1000 in 1% blocking reagent in buffer 1, and the sections were incubated for 1 h at room temperature. Alkaline phosphatase activity was detected using NBT/BCIP substrate (NBT/BCIP 1-step, Pierce Chemical Co., Rockford, IL), the reaction was stopped by immersion in water, and the sections were permanently mounted with GVA Histomount (Zymed).
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Results |
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The pattern of immunostaining in the nonmalignant and malignant
regions of tumor tissue obtained from patients with high grade prostate
cancer using two specific ßA-subunit antibodies is shown
in Fig. 1. Using the ßA
Groome antibody, positive immunoreactivity was observed in basal and
secretory epithelial cells in the nonmalignant regions of tissue (Fig. 1, A and B). Immunoreactivity for ßA-subunit was also
observed in tumor cells from adjacent regions of the tissue containing
poorly differentiated tumor (Fig. 1, D and E). No immunostaining was
observed in either the nonmalignant or malignant regions of control
sections (Fig. 1, C and F). Using the ßA Salk antibody, a
similar pattern of immunostaining was observed; the nonmalignant
epithelium showed positive immunoreactivity that was localized to the
basal and secretory cells (Fig. 1, G and H, indicated by the
arrow), although the intensity of staining was highly
variable. In addition, weaker stromal staining was observed in these
nonmalignant regions of the tissue sections, as demonstrated by the
arrows (Fig. 1, G and H). The malignant epithelial tumor
cells showed consistent and positive staining of localization for
ßA-subunit protein (Fig. 1, J and K). No positive
immunoreactivity was observed with rabbit IgG (Fig. 1, I and L).
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).
Isolated groups of cells in the stroma also expressed ßA
mRNA (shown by the arrows in Fig. 1M). The expression of
ßA-subunit mRNA was detected in the malignant cells of
the adjacent regions of tumor tissue (Fig. 1N). Using the sense
ßA probe, no mRNA expression was detected (Fig. 1, O and
P). ßB-subunit expression and localization
The pattern of expression and localization for
ßB-subunit in nonmalignant and malignant regions of
tissue obtained from patients with high grade prostate cancer is shown
in Fig. 2. Using the ßB
Groome antibody, positive immunoreactivity was primarily localized to
the epithelial compartment of nonmalignant regions of tissue, with more
intense staining of ßB-subunit observed in the basal
cells (Fig. 2, A and B). In adjacent malignant regions,
immunoreactivity for ßB-subunit was localized to the
tumor cells (Fig. 2, D and E), no positive immunostaining was observed
in the control sections (Fig. 2, C and F). Using the ßB
Salk antibody, immunoreactivity was localized to the nonmalignant
epithelial tissue and was predominantly located in the basal, rather
than the secretory, epithelial cells (Fig. 2, G and H). Positive
immunoreactivity for ßB-subunit was also present in the
malignant tumor cells (Fig. 2, J and K). Positive immunoreactivity was
not observed with rabbit IgG (Fig. 2, I and L).
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). mRNA for
ßB-subunit was also expressed in the tumor cells (Fig. 2N). No mRNA expression was detected using the ßB sense
probe in either malignant or nonmalignant regions of the biopsy (Fig. 2, O and P). Localization and expression of follistatin
Using the polyclonal antibody AS 202, follistatin immunoreactivity
was localized to patchy regions of stromal tissue in the nonmalignant
regions of needle biopsies from men with high grade prostate cancer
(Fig. 3, A and B). No immunoreactivity
was observed in the epithelium or in the control sections incubated
with AS 202 preabsorbed with bovine follistatins (Fig. 3C). In
contrast, using the OxB288 antibody, positive immunoreactivity was
localized to the basal cells of the epithelium and was not recorded in
surrounding stromal tissues (Fig. 3, G and H). In the regions of poorly
differentiated tumor, intense positive immunoreactivity was localized
to the malignant epithelial cells using both follistatin antibodies
(Fig. 3, D and E, and J and K). No positive localization was observed
with rabbit IgG (Fig. 3, F and L).
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), consistent with the
immunoreactivity observed with AS 202 (Fig. 3, A and B). Follistatin
mRNA was also expressed in the nonmalignant epithelium, specifically in
the basal cells (Fig. 3M); this site of expression of follistatin mRNA
correlated with the detection of immunoreactivity for follistatin
protein using the OxB288 antibody (Fig. 3, G and H). In the malignant
tumor regions, follistatin mRNA was expressed in the tumor cells (Fig. 3N), which is consistent with the positive immunoreactivity that was
recorded using both follistatin antibodies (Fig. 3, D and E, and J and
K). mRNA expression was not detected using the follistatin sense
riboprobe (Fig. 3, O and P). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In the nonmalignant regions of the tissue sections from men with high grade prostate cancer, activin subunits ßA and ßB were detected in the epithelium, consistent with our previous observations using tissues from men with BPH (13). Expression of mRNAs for the activin subunits in the epithelium was observed and, therefore, confirmed that the prostatic epithelium is a site of synthesis and expression of activin ligands. Although the expression of activin ßA-subunit mRNA was detected in all basal and secretory cells of the nonmalignant epithelia, the protein was not detected in a uniform pattern using the Salk ßA antibody. This pattern of variable staining, where some glands showed intense staining and others showed none at all, was previously observed using an antibody to dimeric activin A to detect this ligand in tissue sections from men with BPH (13). However, the Groome monoclonal antibody consistently detected ßA-subunit protein in basal and secretory cells of the nonmalignant epithelia, which was identical to the uniform detection of mRNA. The reason why there are differences using the two different ßA antibodies is not known.
In contrast to those of ßA, the expression and synthesis
of ßB were consistently observed to be predominantly
localized to the basal, rather than the secretory, cells. This pattern
of subunit localization was identical using both antibodies and was
consistent with the expression of ßB mRNA. The
predominant synthesis and production of the ßA- and
ßB-subunits in the prostatic epithelial cells provides
these cell types with the capacity to form homo- or heterodimers of the
ß-subunits and, thus, activin A, B, or AB ligands. This laboratory
has previously demonstrated that the nonmalignant epithelium expresses
and produces the inhibin -subunit (13), and therefore, the capacity
to synthesize inhibin A or B must also reside within these cells.
The actions of inhibin and activin ligands on nonmalignant prostatic epithelium have yet to be documented. In normal and hyperplastic acini of human prostate, it has been reported that 70% of proliferating cells are localized within the basal cell layer, which has led to the speculation that basal cells have an important role in the proliferative activity of the epithelium (26). It is noteworthy that the basal cells are the predominant site of synthesis of the activin and inhibin subunits, suggesting that these ligands have the potential to influence renewal of the prostatic epithelium. In the human prostate tumor cell lines LNCaP and DU145, a growth inhibitory response to activin A and B has been reported (18, 27, 28), and it is possible that these ligands have a similar action on the nonmalignant epithelium. Indeed, this might be predicted, as activins are known to have antiproliferative actions in other cells and tissues (5, 6, 7) and to induce apoptosis in certain cell lines (9).
In many instances, the inhibins have effects that oppose those of
activins in some cells and tissues (5, 29). The demonstration that
inhibin -subunits are synthesized by the nonmalignant prostate
suggests that they may regulate the actions of activins on the
nonmalignant epithelium. In contrast, the -subunit is down-regulated
in tissues from men with high grade prostate cancer (Mellor SL,
Richards MG, Pedersen JS, Robertson DM, and Risbridger GP, submitted
for publication), and human prostate tumor cells do not express inhibin
-subunit in vitro (14, 15, 16). The loss of -subunit with
the continued expression of the ß-subunits results in the synthesis
of activins, but not inhibins, in the malignant epithelium. However,
the growth inhibitory actions of activin A and B on some prostatic
tumor cell lines are not consistent with the malignant characteristics
of these cells (18), and a mechanism for conferring resistance to
activin ligands must be considered. By analogy with TGFß, it is
possible that there are inactivating mutations of the activin receptors
(30), but an alternate means of regulating activin action is via
follistatins, i.e. binding proteins with a specific and high
affinity for activins.
Follistatin neutralizes the diverse actions of activins, including apoptosis (9), in various cells and tissues by forming an inactive complex with activin. The results described herein demonstrate the expression and localization of follistatin in tumor cells from men with high grade prostate cancer. The localization of the activin subunits and follistatin to the malignant epithelial cells implies that follistatin neutralizes the bioactivity of the activin ligands in poorly differentiated prostate tumors. This is consistent with the in vitro observation that follistatin neutralizes the growth inhibitory effects of exogenously added activin A to the human prostate tumor cell lines LNCaP and DU145 (18); in contrast, PC3 cells are resistant to exogenously added activin A (18). The only difference between the three prostate tumor cell lines that we have observed is the expression of FS288 mRNA by the activin-resistant PC3 cells, but not by LNCaP or DU145 cells in which the predominant mRNA is FS315 (18). The relative levels and types of follistatin produced by the three prostate tumor cell lines are not known. It has been shown that FS288 has a higher neutralizing activity than FS315 (12), and therefore, it is possible that the presence of FS288 is important in conferring activin resistance, as demonstrated in the PC3 cells. In poorly differentiated tumor tissue, follistatin mRNA and proteins, detected by two different antibodies, localize with activin subunit immunoreactivity, which implies that the tumor cells may be resistant to activin in vivo due to the presence of follistatin.
In the nonmalignant tissues adjacent to the high grade tumors, mRNA expression for follistatin was detected in epithelial basal cells and stroma; using OxB288, FS288 protein was localized predominantly to the basal cells. This suggests that the interplay between activin and follistatin is important in regulating activin’s action on basal cells, but the effect of activins on basal cells is not known. In the nonmalignant stromal tissue, follistatin mRNA is expressed, and protein was localized using AS 202, but not the OxB288 antibody. These results suggest that the follistatin proteins produced in basal and stromal cells are different. Furthermore, in malignant and nonmalignant tissues, these data suggest that the expression of follistatin mRNA results in the production of different follistatin isoforms in specific cell types. The relative levels of follistatin proteins and whether they are bound or unbound to activin may also change in the progression to malignancy. The ability to resolve these issues requires the further development of methods that can discriminate among the different forms of follistatin in bound and unbound forms in situ.
Received May 27, 1997.
Revised July 23, 1997.
Accepted August 1, 1997.
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