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Original Studies |
The National Center for Infertility Research and the Reproductive Endocrine Sciences Center, Department of Medicine, Massachusetts General Hospital (C.K.W., W.F.C., A.L.S.), Boston, Massachusetts 02114; Department of Pathology and Laboratory Medicine, Women and Infants’ Hospital of Rhode Island (G.L-M.), Providence, Rhode Island 02905; and Department of Pathology, University of Southern California (W.Z.), Los Angeles, California 90033
Address all correspondence and requests for reprints to: Corrine K. Welt, Department of Medicine, The National Center for Infertility Research and the Reproductive Endocrine Sciences Center, Massachusetts General Hospital, Boston, Massachusetts 02114.
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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All epithelial ovarian tumors expressed mRNA for the 
, ßA, and
ßB subunits; FS 288 and 315; and the activin type IA, IB, II, and IIB
receptors. ßA mRNA expression, as assessed using semiquantitative
RT-PCR, was 3-fold greater in cultured tumor epithelium than in primary
tumors (band density 0.86 ± 0.17 vs. 0.28 ±
0.09; P < 0.01). In addition, ßA mRNA was
abundantly expressed in normal epithelium in culture (n = 2),
whereas only trace amounts were seen in 2/9 primary epithelial
samples.
Activin protein was secreted by 24/25 primary epithelial ovarian tumors (range 0.2–155.8 ng/mL). In contrast, total inhibin was secreted by only 2/25 (range 0.01–0.92 ng/mL), whereas free FS was not detectable in the medium of any tumor (<0.5 ng/mL). Treatment with activin or FS did not consistently affect cell growth. Measurement of serum activin A in a subset of subjects and in 27 additional subjects with epithelial ovarian carcinoma (n = 33) revealed preoperative activin A levels >3 SD above the mean for pre- and postmenopausal women in 13/33 (39%) subjects.
We conclude that in epithelial ovarian cancer: 1) ßA subunit mRNA is expressed, 2) activin protein is secreted more frequently than inhibin and in greater quantities than FS, 3) ßA subunit mRNA expression is greater in neoplastic and normal epithelium in culture than in the primary tissue, 4) the majority of tumors in culture do not respond to activin or FS treatment with proliferation, and 5) serum activin levels may reflect tumor secretion in some patients. Thus, activin A appears to be available as an autocrine/paracrine factor in epithelial ovarian tumors and may contribute to circulating levels, but its role in tumorigenesis has yet to be defined.
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Inhibin and activin are members of the transforming growth
factor-ß superfamily. Inhibin is composed of an and one of two
ß subunits, ßA (inhibin A) or ßB (inhibin B). Activin is a
related dimeric protein composed of two ß subunits, ßA ßA
(activin A), ßB ßB (activin B), or ßA ßB (activin AB) (6).
Elevated levels of inhibin and its subunits have been detected in
subjects with a variety of gonadal stromal tumors (7, 8, 9), most commonly
granulosa cell tumors (10, 11, 12). In addition, elevated inhibin
levels have been detected in subjects with epithelial ovarian
carcinomas, predominantly mucinous cystadenocarcinomas and mucinous
borderline cystic tumors (13).
There is also evidence that the activin family of proteins plays a key
role in ovarian tumor growth. Deletion of the inhibin subunit in a
transgenic mouse model results in development of gonadal stromal tumors
(14) and elevated circulating activin levels (15), suggesting that loss
of the subunit and/or overproduction of activin results in
tumorigenesis. A recent study of six human epithelial ovarian cancer
cell lines revealed absence of inhibin subunit expression and
secretion, and the presence of the activin type II receptor and ßA
and/or ßB subunit expression and secretion (16). In these ovarian
cancer cell lines and in a cell line derived from a gonadal tumor of an
subunit knockout mouse, activin treatment resulted in increased
proliferation, whereas follistatin (FS), an activin binding and
neutralizing protein (17, 18, 19), decreased proliferation in the cell
lines producing activin (16, 20).
Taken together, these studies suggest that ßA subunit messenger RNA
(mRNA) expression and activin secretion in excess of FS and/or in the
absence of subunit results in increased bioavailable activin, which
may ultimately contribute to growth of epithelial ovarian tumors. To
test this hypothesis, expression of , ßA, and ßB subunits; FS;
and activin type I and II receptor mRNA was examined in 25 primary
epithelial ovarian tumors and in a pure population of epithelium
cultured from tumors (n = 7). Activin A, total inhibin, and FS
protein secretion was measured from primary epithelial ovarian tumors
in vitro. mRNA expression and protein secretion from primary
tumors were compared with results derived from the epithelium of 9
normal ovaries and normal ovarian epithelium in culture (n = 2).
Circulating activin A levels were also measured in a subset of these
patients and in 27 additional subjects with epithelial ovarian
carcinoma (n = 33). The resulting demonstration of ßA subunit
mRNA expression and activin A protein secretion in epithelial ovarian
tumors but not normal epithelium, the increase in ßA subunit
expression in neoplastic and normal epithelium propagating in culture,
along with the elevated serum activin A levels in a subset of subjects
with epithelial ovarian carcinoma suggest that activin may be a factor
contributing to tumorigenesis in these patients.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Epithelial ovarian tumors.
Epithelial tumor tissue (
1 g)
was removed by the pathologist from a nonnecrotic portion of the
surgical specimen (n = 25). Epithelial tumor subtypes included 1
borderline serous, 13 serous, 4 endometrioid, 1 borderline mucinous, 3
mucinous, and 3 clear cell carcinomas. Tissue was placed in sterile
Dulbecco’s PBS and transported, on ice, to the laboratory within
18 h (usually <4 h) of the surgery. Tissue was flash frozen and
stored at -80 C for later mRNA extraction (see below). In addition,
0.1–0.4 g of tissue was processed using the explant method (21) in 10
mL RPMI-1640, supplemented with 10% FCS, 2 mM
L-glutamine and antibiotics (100 IU/mL penicillin and 100
µg/mL streptomycin sulfate). The tissue was incubated at 37 C in a
5% carbon dioxide atmosphere for 1 week to allow accumulation of
sufficient hormone for detection while still reflecting secretion of
the primary tumor. Medium was collected, centrifuged to remove tissue,
and stored at -20 C until assayed.
Normal epithelium. Normal ovarian surface epithelium was obtained from 11 control subjects who underwent oophorectomy for a nonovarian cause. Epithelium was scraped from the ovarian surface using a disposable cell scraper (22) and placed in sterile Dulbecco’s PBS on ice. Cells were immediately centrifuged, and the resulting pellet resuspended in 1 mL of Trizol (Gibco BRL, Grand Island, NY) for subsequent mRNA extraction (n = 9) or in RPMI-1640 medium supplemented as above for protein analysis (n = 2).
The procedure for collection of all specimens was approved by the Subcommittee on Human Studies at the Massachusetts General Hospital.
Culture of primary tumors and normal epithelium
Cells remaining after removal of medium for hormone assay were maintained in culture by changing medium weekly, subculturing after cells reached approximately 80% confluence (1 week to 1 month), and removing fibroblasts using selective trypsin treatment (21). To assure that RNA was extracted from a pure population of epithelial tumor cells, the identity of cells in culture was confirmed by positive immunohistochemical staining using mouse antiserum against human cytokeratin (AE1/AE3) and negative staining using mouse antiserum against human desmin (D33) (Dako Corp., Carpinteria, CA). If subculture of cells for immunohistochemistry was impossible because of inadequate growth, morphology was assessed using previously described methods (23, 24). In general, <10% of the cells were nonepithelial by morphology. Because of the failure of many epithelial carcinomas to grow in culture (21) and the slow growth of others, a total of six tumor cultures were used after 3 weeks in proliferation studies and seven cultures were used after 1 month for mRNA extraction and RT-PCR.
Collection of serum samples
After obtaining written informed consent, excess serum or plasma was obtained from preoperative surgical testing and at least 48 h postoperatively in 6 subjects, age 51–75 yr, undergoing surgery for epithelial ovarian cancer. Tumor tissue from the same subject was analyzed as described below. An additional 27 preoperative serum samples and 9 paired postoperative samples were obtained after CA-125 testing from a separate group of subjects with epithelial ovarian cancer, age 40–80 yr. Control subjects included 8 healthy postmenopausal and castrate women, age 51–64 yr, and 6 premenopausal women, age 26–38 yr, with normal menstrual cycles as described previously (25). The blood sample was obtained on the day of menses for premenopausal women to reflect the peak activin A levels in the menstrual cycle (26). The procedures were approved by the Subcommittee on Human Studies at the Massachusetts General Hospital and at Women and Infants’ Hospital of Rhode Island.
Protein measurement
Total inhibin RIA (Monash RIA).
Total inhibin was measured
using the Monash RIA with antiserum 1989 provided by Dr. G. Bialy,
Contraceptive Development Branch, NICHD, as described previously (16).
The inter- and intraassay coefficients of variation were 12% and 10%
respectively. The assay sensitivity was 75 pg/mL. This assay recognizes
monomeric forms of -inhibin as well as dimeric inhibin (27) via an
epitope at the C terminus of the mature subunit (aa 93–108)
(28).
Dimeric inhibin A two-site enzyme-linked immunosorbent assay (ELISA). Dimeric inhibin A was measured in samples with elevated total inhibin using a commercially available ELISA (Serotec Ltd., Oxford, England), as previously described (29). The intraassay coefficient of variation was 9.0%, and the interassay coefficient of variation was 6.8%. The assay sensitivity was 1 pg/mL, and cross-reactivity with inhibin B and activin A was <0.5%.
Activin A ELISA. Total activin A was measured in a two-site solid-phase assay, using a monoclonal antibody against the ßA subunit of inhibin (E4; aa 82–114; 250 ng/well) (Serotec) for both capture and detection according to previously published methods, except where specified (30). Human recombinant activin A, purified in our laboratory from medium conditioned by 293 cells expressing the ßA subunit and calibrated against human recombinant activin A (Genentech Research Reagents Program, South San Francisco, CA), was used as a standard. Sample medium from primary tumors was concentrated approximately 10-fold in a 10,000 mol wt cutoff centricon (Amicon, Beverly, MA). Activin was dissociated from binding proteins by treatment with an equal volume (125 µL) of 8% SDS and incubation at 95 C for 10 min. Assays were developed using streptavidin-alkaline phosphatase (diluted 1:2000) and p-nitrophenyl phosphate. The interassay coefficient of variation was 16% and the intraassay coefficient of variation was 12%. The sensitivity of this assay was 5 ng/mL. Inhibin A cross-reactivity was <0.1%.
Serum activin A was measured using the same assay after it had become commercially available (Serotec) (30), because of the greater sensitivity needed for serum measurements. The assay procedure differed from the previous in that an Ampak substrate kit (Dako Diagnostics, Cambridgeshire, UK), including addition of 1 µL 1 M MgCl2/mL substrate, was used to develop the assay. The assay sensitivity was 80 pg/mL, the value of the lowest standard. The inter- and intraassay coefficients of variation were <7%. There was no cross-reactivity reported with inhibin A, FS, activin B, or inhibin B.
Free FS two-site monoclonal antibody immunoradiometric
assay.
Free FS was measured in a two-site immunoradiometric assay
using two monoclonal antibodies to nonoverlapping epitopes as
previously described (31). The intra- and interassay coefficients of
variation were between 2.7–4.9% and 7.8–11.7%, respectively. The
assay detection limit was 0.5 ng/mL. No cross-reactivity was observed
with inhibin A, activin A, or 2 macroglobulin (31).
Total FS RIA.
Because FS-activin complexes would not be
detected by the free FS assay, tumor media was also analyzed using a
total FS assay that utilizes a polyclonal antibody to human FS 288
(32). The intra- and interassay coefficients of variation were less
than 8% and 11%, respectively. The assay detection limit was 3.2
ng/mL, and no cross-reactivity was demonstrated with inhibin A, activin
A, or 2 macroglobulin.
RNA extraction
Epithelial ovarian tumors. Frozen tumor sample (0.1–0.4 g) was homogenized in 4 mL Trizol (Gibco BRL) using a mechanical tissue homogenizer (PowerGen 125, Fischer Scientific, Pittsburgh, PA), and mRNA extracted according to the manufacturer’s instruction. Total RNA (25–50 µg) was treated with 5 U deoxyribonuclease I, Amp Grade (Gibco BRL) in a 50-µL reaction volume, and the resulting sample reextracted using Trizol. The RNA concentration at each step was determined using spectrophotometric analysis at 260 nm.
Normal epithelium. mRNA was extracted from normal epithelium using Trizol, as above, after addition of 10–20 µg Escherichia coli transfer RNA (tRNA) to facilitate maximal mRNA recovery. Normal epithelium was not subject to DNase treatment.
Normal epithelium and epithelial ovarian tumors in culture. RNA from tumor and normal epithelium in culture was extracted with Trizol after 1 month in culture and treated (1 µg) with 1 U DNase I, as above, in a 10-µL reaction volume.
RT-PCR analysis
Complementary DNA (cDNA) was obtained by RT at 42 C for 45 min
in a 20-µL reaction mixture containing 1 µg RNA, 0.25
mM of each deoxynucleotide triphosphate, 5 µM
oligo deoxythymidine, 200 U Superscript II (Gibco) reverse
transcriptase, and 18 U RNase inhibitor (Promega, Madison, WI). The
cDNA from three separate RT reactions was pooled and subjected to PCR
for ; ßA; ßB; FS (315 and 288); activin type IA, IB, II, and IIB
receptors; and ß-actin targets.For normal epithelium, however, the
amount of mRNA could not be quantified because of the addition of tRNA
to the extraction, and 5–10 µg of tRNA + epithelial RNA was added to
each reaction. The PCR reaction was performed in a volume of 25 µL
containing 0.2 mM of each deoxynucleotide triphosphate, 1
µM 5' and 3' primers, 1 unit Taq polymerase
(Promega), and 1 µCi nucleotide triphosphate
([32P]deoxycytidine triphosphate), to which 5 µL of the
RT reaction was added. All primers crossed at least one intron to
assure that the band resulting from PCR analysis was because of mRNA,
except in the case of ßB (see Table 1
).
For this primer product, a control tube was included for each sample
with no added RT enzyme, resulting in no detectable band in any sample.
In addition, the primer used to detect FS was designed to cross intron
5, which is alternately spliced for transcription of FS 288. Thus,
using one primer set, bands of two sizes could be detected for FS, and
were shown by Southern blot and sequencing to represent FS 288
(835-kilobase band) and 315 (531-kilobase band) (33).
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, ßA, and ßB targets were
inconsistently detected in some samples. Pretreatment of the cDNA
before PCR using 3 U RNase at 37 C for 20 min resulted in consistent
detection and greater sensitivity. The tumor samples were reprocessed
using this method, and results analyzed separately for , ßA, and
ßB. All samples were overlaid with mineral oil, and amplification achieved using a thermal cycler (DNA Thermal Cycler, Perkin Elmer, Norwalk, CT). The amplification profile involved preincubation at 94 C for 5 min, denaturation at 94 C for 1 min, and primer annealing at a temperature decreasing from 65 C to 56 C by 1 min each cycle for 10 cycles, then 55 C for 1 min for an additional 30 cycles and extension at 72 C for 1 min for all cycles. Using undiluted cDNA, PCR amplification was maximal under these conditions, ensuring detection in tissues with low expression.
Five microliters of the PCR reaction was electrophoresed in a 5% polyacrylamide gel in Tris-borate-EDTA buffer. Autoradiography was carried out for 1–2 h at room temperature, and each band scanned on a densitometer with automatic background subtraction.
Based on the consistent and qualitatively greater signal from the ßA
PCR band (relative to ß-actin diluted 104) in cultured
samples compared with primary tumors (Fig. 1A), we developed semiquantitative
conditions to compare ßA expression between the seven tumors in
culture and seven paired primary tumor samples. PCR conditions were
optimized so that the reaction was in the exponential phase when cDNA
(after RT) was diluted 1:100 for ßA and 1:10,000 for ß-actin, and
amplification carried out for 30 cycles, as described above. Both
targets were run for all samples in the same reaction under identical
conditions. The absorption of the resulting ßA band was normalized to
the absorption of the band for ß-actin.
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Thymidine incorporation assay
Cells cultured from tumor and normal epithelium were harvested at approximately 80% confluence and plated in a 24-well plate (1 x 104 cells/well) in RPMI medium supplemented with 10% FBS. At 24, 48, and 96 h, medium was changed to control medium (RPMI with 10% FBS) or control medium supplemented with recombinant human activin A 100 ng/mL, recombinant human FS 288 100 ng/mL (National Hormone Pituitary Program, NICHD) or recombinant human epidermal growth factor (EGF) 10 ng/mL (Sigma Chemical Co., St. Louis, MO). Doses of activin and FS were selected based on their ability to maximally stimulate or inhibit growth in a study of ovarian tumor cell lines (16). At 96 h, 1 µCi [3H]thymidine (6.7 Ci/mmol; NEN, Boston, MA) was added to the medium, and cells incubated for 18–24 h. DNA was subsequently isolated using 10% trichloroacetic acid and solubilized in 1 N NaOH. Each experiment was performed in duplicate.
Statistical analysis
Comparison between the normalized absorbance of PCR bands for ßA in primary and cultured tumors was made using Student’s t test. Correlation between medium or serum activin A or inhibin and tumor subtype was evaluated using ANOVA and a Tukey honest significant difference test for post-hoc comparisons. In addition, a comparison of pre- and postoperative activin A levels was made using a Student’s t test. A P value of <0.05 was considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Epithelial ovarian tumors.
All epithelial ovarian tumors
expressed mRNA for , ßA, and ßB subunits; FS 288 and 315; and
the activin type IA, IB, II, and IIB receptors. ßA mRNA expression in
tumors after 1 month in culture (n = 7) was 3-fold greater than
that of the matched primary tumors (0.86 ± 0.17 vs.
0.28 ± 0.09; P < 0.01) (Fig. 1B
).
Normal epithelium.
Normal epithelium expressed mRNA for FS 288
and 315, and the activin type IA, IB, II, and IIB receptors (Table 2, 1–8). There was detectable
subunit mRNA expression in 3/8 samples but barely detectable ßA
subunit mRNA expression in only 2/8 samples. After 1 month in culture,
however, subunit mRNA was undetectable, whereas ßA was abundantly
expressed in the two samples.
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, ßA, and ßB subunits in
all tumors (n = 25) and tumors in culture (n = 3). In one
additional normal epithelial sample, subunit but not ßA or ßB
was detectable (Table 2, 9), consistent with the results above. Protein secretion
Epithelial ovarian tumors.
Activin protein was secreted by
24/25 (96%) primary tumors (range 0.2–155.8 ng/mL). Total inhibin was
secreted by only 2/25 (8%) tumors. These 2 tumors belonged to the
group of 4 tumors of mucinous subtype; a borderline mucinous
cystadenoma (10 pg/mL) and a low-grade mucinous cystadenocarcinoma (920
pg/mL), which also secreted dimeric inhibin A (118 pg/mL). FS was
secreted by 10/13 (77%) tumors (range 0.5–16 ng/mL), however, free FS
was not detectable in the medium of any tumor (0/25; Fig. 2). Activin A secretion correlated
significantly with tumor type (P < 0.05); mucinous
tumors secreted significantly higher levels of activin A than serous
tumors, although the numbers were small, and the ranges overlapped
(102.1 ± 32.8 vs. 31.9 ± 10.9 ng/mL; Fig. 3A). Mucinous tumors also secreted
significantly higher levels of total inhibin than serous tumors
(0.23 ± 0.23 vs. 0 ng/mL; P < 0.05),
although no overall correlation was seen between total inhibin and
tumor subtype. There was no correlation between inhibin or activin A
secretion and tumor grade.
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Activin A levels in serum.
Activin A levels in pre- and
postmenopausal subjects with epithelial ovarian carcinoma were compared
with values in normal postmenopausal women (0.65 ± 0.18 ng/mL)
and normal premenopausal women on the day of menses (0.66 ± 0.12
ng/mL). Because the activin A levels were not significantly different
in pre- and postmenopausal controls, the groups were pooled, and an
elevated activin A level was defined as >3 SD above the
mean for all normal subjects (1.10 ng/mL). Serum activin A levels were
elevated in 13/33 subjects (39%), with the majority of elevated
activin levels in subjects having the serous subtype (Fig. 3B). The
average activin A level in subjects with epithelial tumors (1.13
± 0.66 ng/mL) and in the subset of subjects with serous tumors
(1.33 ± 0.17 ng/mL) and undifferentiated tumors (1.45 ±
0.76 ng/mL) was significantly higher than in controls (0.65 ±
0.15 ng/mL; P < 0.05 for all groups). In the subset of
subjects with elevated activin A levels in whom both pre- and
postoperative serum samples were available, there was a slight decrease
in activin A postoperatively (1.53 ± 0.16 vs.
1.05 ± 0.20 ng/mL), although this result did not reach
significance (Fig. 4). In the 6 subjects
in whom tumor tissue was also available, there was no correlation
between circulating activin A levels and secretion by the tumor
in vitro. There was also no correlation between circulating
activin A level and tumor subtype.
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In tumor epithelium cultures, activin treatment (100 ng/mL) resulted in increased proliferation in 1/6 tumors, whereas FS treatment (100 ng/mL) increased proliferation in 2/6 tumors. EGF treatment (10 ng/mL), used as a positive control, resulted in a significant increase in proliferation. In normal epithelium cultures (n = 2), EGF treatment resulted in a significant increase in proliferation, whereas no consistent increase or decrease was seen with activin or FS treatment.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, ßA, and ßB subunit; FS; and activin type I and II
receptor mRNA expression and activin A secretion by virtually all of
the epithelial ovarian tumors examined. Further, activin A is secreted
in 10- to 20-fold excess over FS, the protein that binds and
neutralizes activin’s action (17, 18, 19), whereas total inhibin is
secreted by only a subset of mucinous tumors, a finding consistent with
previous immunohistochemical studies (38). In contrast, normal ovarian
epithelium expresses little ßA subunit mRNA and secretes no
detectable activin A but rather favors subunit production. Taken
together, these results demonstrate that epithelial ovarian tumors
produce bioavailable activin, and support the possibility that activin
A may play a role in tumor development and/or growth. Previous studies using normal human granulosa cells (39, 40) and a gonadal stromal tumor cell line (20), and our own studies using epithelial carcinoma cell lines (16), indicate that activin increases cellular proliferation, whereas FS blocks this effect. We were unable, however, to demonstrate a consistent activin effect on proliferation in normal or neoplastic ovarian epithelium in this study, perhaps because of: 1) absence of a necessary cofactor or growth factor secreted by the stroma and required for activin action; 2) loss of contact with other epithelial cells after subculturing; and/or 3) the inherent ovarian epithelial cell senescence after passaging in culture (41). Our data does, however, reveal greater ßA subunit mRNA expression in normal and neoplastic ovarian epithelial cells in culture. This increase may be because of up-regulation of gene expression by factors involved in cell proliferation, or absence of inhibitory factors secreted by the surrounding stroma and lost in culture. Taken together, the evidence suggests that activin production increases in proliferating ovarian epithelium and may act in an autocrine or paracrine manner to stimulate growth. Thus, if the frequent cell division required for epithelial repair results in a genetic mutation and transformation, activin may act in a permissive manner to promote neoplastic cell growth. Alternatively, loss of an inhibitory growth factor such as tranforming growth factor-ß (42), or a mutation resulting in dysregulation of the control or response mechanism of activin could lead directly to uncontrolled growth.
Despite inhibin subunit mRNA expression by all tumors, secretion
was demonstrated by only 2 mucinous tumors. This discrepancy suggests:
1) a translational block; 2) mutation or alternate processing of the
subunit rendering it undetectable in the Monash assay; and/or 3)
secretion at levels below the assay detection limit. In contrast,
detectable inhibin was secreted by 2/2 normal epithelial cultures.
These findings suggest the possibility that subunit could act as a
tumor suppressor as was initially described in the subunit knockout
mouse (14). Rather than tumorigenesis resulting from deletion of a
tumor suppressor gene, however, subunit may act as a tumor
suppressor protein by dimerizing with ß subunit, thus preventing
activin formation and decreasing proliferative potential. The
difference in tumor subtypes in the mouse (gonadal stromal tumors) and
human (epithelial tumors) could be related to the potentially different
tumor suppressor mechanisms or more likely to the differing ovarian
biology in the two species, because epithelial tumors are rare in the
mouse (43).
Activin A levels are increased in the serum of subjects with epithelial
tumors as demonstrated by an elevated mean activin A level for all
subjects and elevated levels in a significant number of subjects
compared to controls. The majority of elevated levels are accounted for
by subjects with serous and undifferentiated tumors and the subset of
mixed tumors containing both serous and endometrioid components.
Therefore, measurement of activin A levels in serum may provide
information additive to serum inhibin levels, which are elevated in
many patients with mucinous but not serous tumors (12, 13). In contrast
to these serum findings, however, mucinous tumors secrete the highest
level of activin A in vitro and are the only epithelial
tumor subtype that demonstrates positive staining for immunoreactive
activin (38). The small number of mucinous tumors examined or the
unavailability of paired tissue and serum samples may account for the
discrepancy between serum and in vitro data, although we
cannot exclude the possibility that mucinous tumors secrete an activin
binding protein that sequesters activin locally. Based on the large
number of subjects with elevated activin A levels and the apparent
postoperative decrease in subjects with preoperative serum elevation,
it appears that tumor production of activin A may contribute to serum
levels. Thus, activin A may potentially serve as a tumor marker for
serous and undifferentiated tumors. Further investigation using a
larger number of subjects is needed to document a postoperative fall in
activin A and to correlate in vitro activin A production
with serum levels.
In summary, the data provided here clearly demonstrate activin subunit and receptor mRNA expression and secretion of activin A in excess of the activin binding protein FS in epithelial ovarian cancers but not normal epithelium. Furthermore, circulating activin A levels are elevated in a subset of epithelial tumors. Therefore, further investigation of activin A as both a potential growth factor and a tumor marker is warranted.
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Acknowledgments |
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Footnotes |
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Received March 5, 1997.
Revised May 20, 1997.
Revised July 8, 1997.
Accepted July 15, 1997.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and ß subunits of
inhibin/activin as markers for gonadal sex-cord stromal
differentiation. Int J Gyn Pathol. in press.
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F. M. Reis, L. Cobellis, L. C. Tameirao, G. Anania, S. Luisi, I. S. B. Silva, W. Gioffre, A. M. di Blasio, and F. Petraglia Serum and Tissue Expression of Activin A in Postmenopausal Women with Breast Cancer J. Clin. Endocrinol. Metab., May 1, 2002; 87(5): 2277 - 2282. [Abstract] [Full Text] [PDF]
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G. P. Risbridger, J. F. Schmitt, and D. M. Robertson Activins and Inhibins in Endocrine and Other Tumors Endocr. Rev., December 1, 2001; 22(6): 836 - 858. [Abstract] [Full Text] [PDF]
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T. Fujiwara, Y. Sidis, C. Welt, G. Lambert-Messerlian, J. Fox, A. Taylor, and A. Schneyer Dynamics of Inhibin Subunit and Follistatin mRNA during Development of Normal and Polycystic Ovary Syndrome Follicles J. Clin. Endocrinol. Metab., September 1, 2001; 86(9): 4206 - 4215. [Abstract] [Full Text] [PDF]
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K.-C. Choi, S. K. Kang, C.-J. Tai, N. Auersperg, and P. C. K. Leung The Regulation of Apoptosis by Activin and Transforming Growth Factor-{beta} in Early Neoplastic and Tumorigenic Ovarian Surface Epithelium J. Clin. Endocrinol. Metab., May 1, 2001; 86(5): 2125 - 2135. [Abstract] [Full Text]
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N. Auersperg, A. S. T. Wong, K.-C. Choi, S. K. Kang, and P. C. K. Leung Ovarian Surface Epithelium: Biology, Endocrinology, and Pathology Endocr. Rev., April 1, 2001; 22(2): 255 - 288. [Abstract] [Full Text]
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 F. M. Reis, A. Faletti, S. Luisi, G. Bifulco, S. Cauci, F. Quadrifoglio, J. Dor, and F. Petraglia High concentrations of inhibin A and inhibin B in ovarian serous cystadenoma: relationship with oestradiol and nitric oxide metabolites Mol. Hum. Reprod., December 1, 2000; 6(12): 1079 - 1083. [Abstract] [Full Text] [PDF]
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 T. Minegishi, T. Kameda, T. Hirakawa, K. Abe, M. Tano, and Y. Ibuki Expression of Gonadotropin and Activin Receptor Messenger Ribonucleic Acid in Human Ovarian Epithelial Neoplasms Clin. Cancer Res., July 1, 2000; 6(7): 2764 - 2770. [Abstract] [Full Text]
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 J. L. Juengel, L. D. Quirke, D. J. Tisdall, P. Smith, N. L. Hudson, and K. P. McNatty Gene Expression in Abnormal Ovarian Structures of Ewes Homozygous for the Inverdale Prolificacy Gene Biol Reprod, June 1, 2000; 62(6): 1467 - 1478. [Abstract] [Full Text]
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C. K. Welt, D. J. McNicholl, A. E. Taylor, and J. E. Hall Female Reproductive Aging Is Marked by Decreased Secretion of Dimeric Inhibin J. Clin. Endocrinol. Metab., January 1, 1999; 84(1): 105 - 111. [Abstract] [Full Text]
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Y. Sidis,, T. Fujiwara,, L. Leykin,, K. Isaacson,, T. Toth,, and A. L. Schneyer Characterization of Inhibin/Activin Subunit, Activin Receptor, and Follistatin Messenger Ribonucleic Acid in Human and Mouse Oocytes: Evidence for Activin's Paracrine Signaling from Granulosa Cells to Oocytes Biol Reprod, October 1, 1998; 59(4): 807 - 812. [Abstract] [Full Text]
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