Department of Pathology and Laboratory Medicine (S.L.A., L.R.)
Mount Sinai Hospital and Department of Laboratory Medicine and
Pathobiology University of Toronto Toronto, Ontario, Canada M5G
2M9
Department of Physiology and Biophysics (P.R.M.,
A.W.L.) Dalhousie University Halifax, Nova Scotia, Canada B3H
4H7
Department of Medicine (S.E.) Mount Sinai
Hospital and University of Toronto Toronto, Ontario, Canada M5G
2M9
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The biology of these tumors provides an opportunity to identify novel
mechanisms of dysregulated cell proliferation. Although they are
monoclonal in origin, the molecular events responsible for pituitary
cell transformation are not known (2). Their relatively indolent
behavior is consistent with lack of somatic mutations that are
characteristic of other human malignancies. Because these tumors form
an integral part of the multiple endocrine neoplasia type 1 (MEN-1)
syndrome, the MEN-1 tumor suppressor gene was a candidate
gene responsible for these tumors. However, loss of heterozygosity with
mutations and down- regulation of the menin gene are
rare in the more common sporadic tumors. Activating mutations of the
Gs protein have been implicated in a minority of GH-producing
adenomas. Hypothalamic stimulation has been implicated and appears to
be important in promotion of tumor growth but not cell transformation.
Mice that overexpress GH-releasing hormone (GHRH) and mice that lack
dopaminergic D2 receptors that mediate inhibition of PRL secretion
develop target adenohypophysial cell hyperplasia that then predisposes
to subsequent neoplastic transformation of pituitary tumors.
However, these models differ from the human disease, since patients
with pituitary tumors do not exhibit underlying hyperplasia (2).
Growth factors have been implicated in pituitary tumorigenesis. The pituitary is the site of synthesis and the target of several growth factors that modulate hormone production and are believed to regulate, in part, pituitary cell growth (3, 4). At least 19 members of the fibroblast growth factor (FGF) family have been described with variable mitogenic, angiogenic, and hormone regulatory functions (5). Fibroblast growth factor-2 [FGF-2 or basic FGF (bFGF)] was, in fact, originally isolated from bovine pituitary (6) and is differentially expressed by pituitary adenoma cells with higher levels noted in the more aggressive tumors (7). FGF-2 is known to stimulate vascular endothelial growth factor (VEGF) expression as a mechanism of inducing angiogenesis, suggesting a possible indirect mechanism of tumorigenesis (8).
The regulation of FGF-2 gene expression is poorly understood. However, one possible mechanism involves posttranscriptional regulation of the FGF-2 mRNA by interaction with an FGF antisense RNA. A 1.5-kb FGF-2 antisense (GFG) RNA complementary to the third exon and 3'-untranslated region (UTR) of FGF-2 mRNA has been indirectly implicated in FGF-2 mRNA editing and stability (reviewed in Ref. 9). FGF antisense gene expression has been identified in a number of species including avian, rodent, and human (10, 11, 12). Indirect evidence supports the hypothesis that the antisense RNA may regulate FGF-2 in these species. Steady-state levels of the antisense RNA are inversely related to the level of FGF-2 mRNA during embryonic development and, in a variety of tumor cell lines (11, 13, 14, 15, 16). We recently demonstrated that expression of the FGF antisense RNA in rat C6 glioma cells suppressed FGF-2 expression and inhibited cell proliferation (17).
In addition to its putative role as a regulatory RNA, the antisense transcript also encodes a translated 35 kDa protein (GFG) with homology to the MutT family of antimutator nucleotide hydrolases (NTPases) (14). Although GFG can partially complement function in MutT-deficient Escherichia coli (18), the physiological function of GFG in mammalian cells is unknown.
We hypothesized that human GFG may play a role in modulating the proliferative and hormone-regulatory actions of FGF in the pituitary and may be implicated in the unique biological behavior of pituitary adenomas.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
Using immunohistochemistry, nontumorous pituitary exhibited cytoplasmic
GFG protein throughout the gland with variable levels detectable in
individual cells (Fig. 3a). Localization
did not correlate with hormone content; double staining for pituitary
hormones showed colocalization of GFG with each of the pituitary
hormones (data not shown). Pituitary tumors exhibited variable
intensity of diffuse cytoplasmic staining for GFG (Fig. 3a
) that
correlated with mRNA expression but not with tumor type. More
aggressive and recurrent adenomas lacked detectable GFG
immunoreactivity.
|
GFG Inhibits Cell Proliferation in Vitro
To determine whether GFG is of oncogenic significance, we
produced stably transfected lines of GH4 cells expressing GFG and
measured cellular proliferation by
[3H]thymidine incorporation. Compared with
control cells transfected with empty vector or the untranslated splice
variant, cells expressing GFG exhibited reduced proliferative activity,
ranging from 65 to 80% of control. Among 12 clones that expressed GFG,
the degree of inhibition of thymidine incorporation was
proportional to levels of GFG protein expression. The highest GFG
stably expressing clone demonstrated the greatest reduction of
proliferative activity (Fig. 4A).
Inhibition of proliferation was further confirmed by proliferating cell
nuclear antigen (PCNA) labeling indices that were 30% in control cells
and were reduced to 812% in 12 stably transfected cell lines (Fig. 4A
).
|
Flow cytometric evaluation of cell cycle in GH4C1 cells showed at least
a 50% reduction in S phase fractions in the cell lines stably
transfected with GFG (Fig. 4B). Again, the degree of reduction of the
proportion of cells in S phase was proportional to levels of GFG
protein expression in the various clones.
To exclude the possibility of clonal selection as an independent cause
of reduced cell proliferation, we performed the experiments on cells
transiently transfected with the GFG expression vector, the empty
vector, or the untranslated splice variant. Again there was persistent
reduction of thymidine incorporation (<75% of control) as well as a
marked reduction (>50%) in S phase fraction in cells transiently
transfected with GFG compared with control cells expressing empty
vector or the untranslated splice variant. ß-Galactosidase staining
revealed a transfection efficiency of more than 50%, and trypan blue
exclusion by more than 90% of cells exclude the possibility that the
reduction in cell proliferation was attributed to or associated with
nonspecific toxicity. Western blotting confirmed expression of
the transfected protein (Fig. 5A).
|
Conversely, to determine whether FGF regulates GFG translation, we exposed transfected cells expressing GFG to FGF-2. There was no change in GFG protein in the cells exposed to FGF stimulation, excluding FGF modulation of GFG directly.
GFG Regulates Pituitary Hormones
Since FGF is known to stimulate expression of the PRL gene in
GH4C1 cells, we used this model to determine whether GFG plays a role
in hormone production. Compared with cells stably or transiently
transfected with empty vector or the untranslated splice variant, those
expressing GFG showed significantly enhanced PRL-luciferase activity
(Fig. 6, A and C) (P <
0.05) and had markedly increased PRL immunoreactivity in cell lysates
and media as determined by Western blotting (Fig. 6B
). GFG transfection
resulted in no increase in GH-luciferase activity, which was even
slightly reduced (Fig. 6
, A and C). Total GH immunoreactivity as
determined by Western blotting and densitometry was unchanged; however,
GFG expression was associated with an increase of the 22-kDa form over
the 20 kDa form of GH (Fig. 6B
).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In Xenopus laevis oocytes, a 1.5-kb FGF-2 antisense RNA complementary to the third exon and 3'-UTR of the FGF-2 mRNA has been suggested to play a role in regulating FGF-2 mRNA editing and stability (26). In addition, however, the antisense RNA encodes a distinct protein of poorly defined function. We have previously shown that this predicted protein, with homology to the MutT family of nucleotide hydrolases, is expressed in non-CNS tissue in the postnatal period (11). Here we show, for the first time, that the human pituitary expresses the FGF-AS mRNA and that the predicted protein GFG product is efficiently translated and can be detected by Western blotting and immunohistochemistry. Moreover, we demonstrate that this protein is localized in the cytoplasm in pituitary cells. This is in contrast to the predominantly nuclear localization of GFG detected in rat C6 glioma cells. Subcellular distribution of GFG may be determined by alternative splicing, as has been reported for BRCA1 (27) and the human MutY homolog (28). C6 cells predominantly express a 28-kDa isoform of GFG that is derived by alternative splicing to remove exon 2 of the FGF antisense pre-mRNA (17). Alternatively, nuclear vs. cytoplasmic distribution may be a result of lineage-dependent cell-specific factors, as has been reported for p53 (29) and for the homeodomain protein OTX2 (30).
Pituitary-derived FGF-2 has been shown to stimulate replication of
PRL-secreting cells but also may inhibit DNA synthesis in pituitary
adenoma cells (31), suggesting that some forms of the growth factor or
its receptor may act as growth inhibitors. Elevated blood
concentrations of bFGF-like immunoreactivity have been documented in
patients with MEN-1 (32) and in patients with sporadic pituitary
adenomas (7). The FGF-related hst has
been found in transforming DNA of human PRL-secreting tumors (33),
which also facilitates lactotroph proliferation (34). Transgenic mice
expressing bFGF under the control of the GH and the glycoprotein
-subunit promoters developed hyperplasia of several adenohypophysial
cell types but not frank adenomatous changes (35). FGF-2 or homolog
family members have therefore been suggested to play an important
role in pituitary tumor cell replication.
In this study we have found that the normal pituitary and the less aggressive pituitary adenomas expressed relatively more GFG than FGF-2. This may be of functional significance inasmuch as we have demonstrated that GFG expression is associated with diminished cell replication. We also show here that expression of GFG results in restrained pituitary cell growth, an effect supported by diminished thymidine incorporation, PCNA cell labeling, as well as reduced entry into the S-phase of the cell cycle. While we noted that the more aggressive and recurrent adenomas had reduced GFG expression, we did not observe a strict discordant pattern between FGF-2 and GFG expression in primary pituitary adenomas. Moreover, forced expression of GFG in GH4 pituitary cells did not result in parallel reduction in FGF-2 mRNA or protein expression. The inhibition of proliferation was not seen with transfection of a splice variant of GFG that retains the mRNA homology with FGF-2 mRNA but fails to express functional GFG protein. Taken together, these data indicate that GFG plays a direct antiproliferative role in pituitary cell replication that is independent of FGF-2 expression.
FGF-2 and FGF-4 have been shown to induce PRL gene expression (36). In contrast to other systems, however, this FGF effect is independent of Ras or Raf kinase activation. FGF induction of the PRL gene is dependent on mitogen-activated protein (MAP) kinase activity with defined Ets binding sites (36). Antagonism of FGF-2 expression would, therefore, have been expected to result in inhibition of PRL. Instead, we demonstrate that GFG expression results in PRL stimulation, as shown by endogenous PRL levels and transfected PRL-promoter activity. These data further support the notion that GFG expression plays a hormone-regulatory role that is independent of FGF-2 expression. Further studies are underway to determine whether FGF and GFG mediate their effects on the PRL gene through similar signaling cascades.
In contrast to PRL, FGFs have not been shown to regulate the GH gene. In the current study we did not identify stimulation of GH-promoter activity by GFG. Instead, GFG expression was associated with an increase of the 22-kDa isoform relative to the 20-kDa form of GH. Pituitary GH mRNA undergoes alternative splicing into a 20-kDa and 22-kDa isoforms. GFG-induced alteration in the ratio of 22/20 kDa GH isoforms suggests a possible role for GFG in regulating GH mRNA splicing. This finding would suggest that in some systems, GFG may play a role in regulating pituitary gene splicing. It will be particularly interesting to determine whether GFG regulates other non-FGF-related gene splicing. The 20-kDa form of GH has been shown to be a weaker agonist for hPRLR than 22-kDa hGH (37). In the case of GH, the relative increase in the 22-kDa form may serve to enhance responsiveness to PRL action in addition to the direct stimulatory role of GFG on PRL expression.
Our current findings of the expression of the endogenous FGF antisense gene extend our understanding of the role of FGF in modulating the balance of FGF function in the pituitary. The observation that GFG leads to restrained pituitary cell growth coupled with the induction of PRL gene expression are consistent with the common occurrence of small pituitary tumors in patients with hyperprolactinemic disorders.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
mRNA Analysis by RT-PCR
Total RNA was extracted by the guanidinium isothiocyanate
method. One microgram of DNase-treated RNA was used for reverse
transcription. This was performed using 2.5 U/ml of murine leukemia
virus reverse transcriptase, 2.5 mM
MgCl2, 1 mM deoxynucleoside
triphosphate (dNTP), 2.5 mM random hexamers, and 1
U/ml of RNase inhibitor. The integrity of RNA from each sample was
assessed by amplification of the PGK-1 housekeeping gene as previously
described (38). PCR analyses were performed with primers within the
coding human sequences to amplify a 301-bp fragment corresponding to
GFG exons 4 and 5, which is common to all three GFG splice variants
(17), and a 375-bp fragment of the coding region of FGF-2 (7). All
primers were designed to span at least one intron to permit the
exclusion of genomic DNA contamination. PCR conditions were optimized
to ensure product linearity. The identity of all PCR products was
confirmed by Southern blotting hybridization and by sequencing.
Transfection and Plasmid Constructs
Plasmids containing full-length human GFG cDNA or an
untranslated GFG splice variant that lacks exons 2 and 3 but retains
the regions of homology to FGF-2 (17) were prepared by subcloning into
the pcDNA3.1 (Invitrogen, San Diego, CA)
eukaryotic expression vector. The resulting products were subjected to
sequencing for confirmation of sequence fidelity before transfection
with lipofectamine (Life Technologies, Inc., Gaithersburg,
MD) into wild-type GH4C1 cells. Stably transfected cells were isolated
by G418 selection. As inhibition of FGF-2 expression might bias the
selection of transfectant lines in favor of FGF-2 independence, cells
were grown and passaged in the presence and absence of added FGF-2. GFG
expression was confirmed by Western blotting and immunohistochemistry.
To establish transfection efficiency and to allow comparison within and
between experiments, 20 ng/well of pSV-ß-galactosidase control vector
(Promega Corp., Madison WI) was included with each
transient transfection and measured on cell lysates by colorimetric
analysis or in culture by light microscopy.
Chromosomal Localization
The human GFG gene chromosomal localization was determined by
PCR of somatic cell hybrid and radiation hybrid panels by PCR and
hybridization with a human full-length GFG cDNA. Further detailed
localization used a panel of YAC clones for PCR.
Protein Extraction and Cell Fractionation
Total protein was extracted from total cell lysates and media
and quantified. Cell fractionation was performed by the hypotonic/NP-40
lysis method. Briefly, cells were washed in Tris-buffered saline (TBS),
swollen in homogenization buffer [10 mM HEPES, pH 7.9, 10
mM KCl, 0.1 mM EDTA, 0.1 mM
ethylene glycolbis, 1 mM dithiothreitol (DTT), and 0.5
mM phenylmethylsulfonyl fluoride (PMSF)], and vortexed in
homogenization buffer containing 0.6% NP-40. Supernatant containing
cytoplasm and plasma membranes were removed, and the pellet containing
the nuclear fraction was suspended in a resuspension buffer (250
mM Tris, pH 7.8, 60 mM KCl, 1 mM
DTT, and 1 mM PMSF).
Western Blot Analysis
Protein concentrations were determined by the Bio-Rad Laboratories, Inc. (Hercules, CA) protein assay. Equal amounts
of protein (50 µg) from cell lysates or media were solubilized in
2.5x SDS-sample buffer and separated on SDS-8% polyacrylamide gel and
transferred to nitrocellulose. Apparent molecular weights were
determined by comparison with concurrently electrophoresed standards.
GFG, FGF-2, PRL, and GH protein levels were determined using the
following antibodies: polyclonal antisera raised against synthetic
peptides corresponding to the C-terminal epitope of deduced GFG (14)
applied at a dilution of 1:1500; a polyclonal antiserum that recognizes
FGF-2 (Transduction Laboratories, Inc., Lexington, KY)
applied at a dilution of 1:300 (of 100 µg/ml affinity-purified
IgG); and polyclonal antisera to rat PRL or GH (donated by the National
Hormone and Pituitary Program (NHPP), NIDDK, NICHHD, Bethesda, MD)
applied at dilutions of 1:8,000 and 1:50,000, respectively. An actin
control was performed using a monoclonal antibody (Sigma,
St. Louis, MO) at 1:500.
Immunocytochemical Localization of GFG
For immunolocalization, a polyclonal antiserum that recognizes
the MuT domain and another directed against the C-terminal tail of
human GFG (14) was applied at a dilution of 1:300. Primary human
tissues and tumors were fixed in formalin and embedded in paraffin; the
immunolocalization was detected with the streptavidin-biotin-peroxidase
complex technique and 3,3'-diaminobenzidine (DAB) and visualized with a
light microscope. For colocalization with pituitary hormones, double
staining was performed with the following primary antibodies and
antisera: ACTH and GH [polyclonal antisera from DAKO Corp. (Carpinteria, CA) prediluted 1:15 and 1:1,500
respectively); PRL [prediluted monoclonal antibody from Biomeda Corp.
(Foster City, CA)], -subunit of glycoprotein hormones [monoclonal
antibodies from Amac Inc. (Westbrook, ME) and Zymed Laboratories, Inc. (South San Francisco, CA) 1:200 and 1:4]; ß-TSH,
ß-FSH, and ß-LH (monoclonal antibodies from Amac Inc., diluted
1:500, 1:400, and 1:400, respectively). For the double stain to
colocalize GFG and hormone, the streptavidin-biotin-peroxidase method
was used to detect one antigen, and a peroxidase-conjugated secondary
antibody method was used for the other antigen to avoid cross-reaction;
the second chromogen was cobalt DAB. The order and technique of primary
antibody detection were reversed to accurately evaluate cross-reaction
by the primary antibodies. For subcellular localization, transfected
cells were grown on glass coverslips and fixed in 1% formalin in PBS,
and the primary GFG antiserum was localized with fluorescein-tagged
secondary antibody and visualized with a MRC 600 confocal microscope
(Bio-Rad Laboratories, Inc.). The specificity of all
reactions was verified by replacing the primary antibody with normal
rabbit serum and by examining negative controls.
Mitogenic Assay
GH4C1 cells were transfected with GFG, the splice variant, or
empty vector, and cell proliferation was measured by
[3H]thymidine incorporation. Cells were grown
in six-multiwell microtiter plates (5 x 104
cells per well), labeled with 1 Ci/ml
[3H]thymidine for 6 h and collected, and
the amount of trichloroacetic acid (TCA)-precipitable radioactivity
associated with the cells was measured and normalized for total cell
number.
Cell proliferation in transfected cells was also analyzed by PCNA labeling. Cultured cells were collected in pellets, fixed in formalin, and embedded in paraffin. Sections (4 µm-thick) were stained using the PCNA monoclonal antibody (Novocastra, Newcastle-Upon-Tyne, UK) applied at 1:4,000, and detected with the streptavidin-biotin-peroxidase complex technique and DAB. Labeling indices were determined by counting 1,000 cells for each sample, and the labeled cells were expressed as a percentage of total cells. All analyses were performed in triplicate.
Apoptosis and Toxicity Assays
The rate of apoptosis was measured in GH4C1 cells transfected
with GFG, the splice variant or empty vector using DNA nick-end
labeling. Cultured cells were collected in pellets, fixed in formalin,
and embedded in paraffin. Nuclei of tissue sections were stripped of
proteins by incubation with 20 µg/ml proteinase K
(Sigma) for 15 min at room temperature. Endogenous
peroxidase was inactivated with 2%
H2O2 for 5 min. The
sections were immersed in TdT buffer (Oncor, Intergen, Purchase,
NY) (30 mM Trizma base, pH 7.2, 140 mM
sodium cacodylate, 1 mM cobalt chloride). Sections were
then incubated in TdT (30% solution) and biotinylated dUTP in TdT
buffer (Oncor) in a humid atmosphere at 37 C for 60 min. The reaction
was terminated by transferring the slides to TB buffer (300
mM sodium chloride, 30 mM sodium citrate) for
15 min. The sections were blocked with a 2% aqueous solution of human
serum albumin (HSA) for 10 min, rinsed and detected with streptavidin
peroxidase, diluted 1:20 in water, and incubated for 30 min at 37 C and
with DAB for another 30 min at 37 C. Labeling indices were determined
by counting 1,000 cells for each sample and expressed as the percentage
of labeled cells. All analyses were performed in triplicate.
As this assay does not identify nonspecific toxicity, we performed trypan blue exclusion analysis of transfected cells.
Cell Cycle
To identify the cell cycle effects of GFG, cells were analyzed
by fluorescence activated cell sorting (FACS). For cell cycle analysis,
13 x 106 cells were trypsinized, washed
with PBS, and fixed with 80% ethanol for 1 h on ice. The fixed
cells were washed with staining buffer (0.2% Triton X-100 and 1
mM EDTA, pH 8.0, in PBS) and resuspended in the staining
buffer containing 100 µl (10 mg/ml) RNAse A (Sigma) and
50 µl (1 mg/ml) propidium iodide for 1 h. Cell cycle analysis
was done by a FACScan (Becton Dickinson and Co., San Jose,
CA) using the Cellquest Analysis program, and specific S-phase was
analyzed using Modfit DNA Analysis program (Verity Software, Inc.,
Topsham, ME).
Hormonal Regulation
Expression of endogenous PRL and GH was determined by Western
blotting of cell extracts and media from transiently and stably
transfected and control cells as described above and normalized to
actin. Relative concentrations were determined by densitometric
analysis of autoradiographs.
PRL promoter activity was analyzed with reporter constructs pSV2A-rPRL-luc containing the 422-bp fragment of the rPRL promoter or 320 bp of the rGH promoter (kindly provided by Dr. H. Elsholtz, Toronto). To normalize for transfection efficiency variation within and between experiments, 20 ng/well of pSV-ß-galactoside control vector (Promega Corp.) was included with each transfection. The results were normalized to ß-galactosidase activity.
Stimulation of PRL by FGF was analyzed in GH4C1 cells transfected with GFG, the splice variant of GFG, or empty vector. Transfected cells were grown in six-multiwell microtiter plates (5 x 104 cells per well), preincubated for 48 h in serum-free defined media [insulin (5 g/ml), transferrin (5 g/ml)], and then treated with and without FGF-1 (Sigma, 50 ng/ml) and 10 U/ml of heparin in serum free medium for 24 h at 37 C.
Statistical Analyses
Data are expressed as mean ± SEM. Differences
were examined by one-way ANOVA or Students t-test both
with significance level of <0.05.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
FOOTNOTES |
---|
This work was supported by the Medical Research Council of Canada Grants MT-14404 (to S.E.) and MT-14464 (to S.L.A.).
Received for publication November 17, 2000. Revision received December 18, 2000. Accepted for publication January 9, 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|