From the Department of Biology, Villanova University,
Villanova, Pennsylvania 19085 and ¶ Weis Center for Research,
Pennsylvania State University, College of Medicine,
Danville, Pennsylvania 17822
Received for publication, May 15, 2000, and in revised form, October 5, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The immature rat uterus has been extensively used
as an in vivo model system to study the molecular
mechanisms of steroid hormone actions. In this study, we demonstrated
the regulated expression of syndecan-3 in the rat uterus by the steroid
hormone 17 It has been known for some time that ovarian hormones induce
changes in expression of surface proteins on epithelial cells lining
the uterine lumen believed to be crucial for blastocyst reception (1).
Included in this group of modified surface proteins are the heparan
sulfate proteoglycans
(HSPGs).1 HSPGs are nearly
ubiquitous components of mammalian cell membranes that exhibit binding
interactions with a variety of extracellular ligands. Molecular cloning
and analysis has led to identification of multiple forms of
membrane-anchored proteoglycans. Those best characterized are the
membrane intercalated molecules, four of which display significant
homologies in core protein structure and are classified as members of
the same gene family. These related molecules are called the syndecans
and include syndecans-1-4. Syndecans contain a small core protein with
distinct functional domains to which are linked multiple
glycosaminoglycan (GAG) chains, predominantly of the heparan sulfate
variety. Homology of syndecan family members is most striking in the
transmembrane and cytoplasmic domains, showing >50% amino acid
sequence similarity. In contrast, the extracellular ectodomains show
little homology (2). Sequence-specific serine residues in the
ectodomain regions serve as target sites for GAG attachment during
proteoglycan synthesis. Variation in number and position of GAG
attachment sites within the nonhomologous ectodomains leads to
substantial differences in syndecan structure and potentially produces
differential functions for each of the family members.
Precise functions of each syndecan family member have not fully been
determined. Experimental observations indicate involvement of these
compounds in cell adhesion, cell morphogenesis, and regulation of cell
responsiveness to soluble growth-regulatory compounds (3). These
functions are attributed primarily to the glycan moieties. Both basic
fibroblast growth factor (4, 5) and vascular endothelial growth
factor (6) have binding affinities for heparan sulfate, and evidence
indicates that HSPGs function as coreceptors for these compounds at the
cell surface. By virtue of their HSPG associations, binding of growth
factors to specific cell surface receptors is facilitated, thereby
enhancing effective signal transduction in target cells. Heparan
sulfate GAG chains also provide cell membrane binding sites for
extracellular matrix adhesion proteins, including fibronectin (7, 8)
and laminin (9). Such binding interactions may facilitate or reinforce attachment of matrix proteins to their specific cell surface integrin receptors. Evidence that cell surface HSPGs have a role in focal adhesion formation in fibroblast and Chinese hamster ovary cells (10)
and the specific localization of syndecan-4 to focal adhesions in a
variety of cell types (11, 12) further supports syndecan facilitation
of organized cell-matrix interactions. Syndecan-3 has been implicated
as an important matrix adhesion molecule in regulated neurite outgrowth
activity during development in the central nervous system (13).
Additional observations that neuronal cell adhesion molecule, a
homophilic cell-cell adhesion protein, exhibits binding affinity for
heparan sulfate also suggest potential HSPG involvement in cell-cell
anchorage (14). A direct assessment of differential binding affinities
of the various membrane proteoglycans has not yet been completed.
Therefore, the functional significance of the diverse forms of syndecan
family members remains unclear.
Given the potential functional variation of the syndecans, it is not
surprising that regulated expression of the family members has been
observed. Although data are incomplete, syndecans appear to be
expressed in tissue-specific patterns, although some crossover is
apparent (for review, see Ref. 15). Syndecan-3, although initially
characterized by its high degree of expression in the nervous system
during early postnatal development (16), is also observed in vascular
smooth muscle cells and stratified epithelia. Regulation of expression
in these various cell types, however, is poorly understood, although
evidence indicates both temporal and spatial changes during a variety
of development and cell differentiation processes. Such changes in
expression often correlate with intervals when cell morphology is
altered and major tissue ultrastructural reorganizations are occurring.
Limited information is available concerning the molecular mechanisms of
syndecan gene regulation. Induction of syndecan-1 expression has been
observed in response to basic fibroblast growth factor in cultured
populations of epithelial cells (17) and in vascular smooth muscle
cells by exposure to platelet-derived growth factor (18). Likewise,
syndecan-4 expression in vascular smooth muscle cells is induced by
basic fibroblast growth factor (19). Ovarian hormones have been shown
to regulate expression of syndecan-1 in mouse uterine tissue (20). In
this study, histochemical analysis to detect binding of anti-syndecan-1
antibodies demonstrated a dramatic change in lumenal epithelial cell
localization during the estrous cycle involving repositioning from a
predominantly apical location during intervals of low hormone secretion
to an exclusively basolateral position during the peak fertility phase. Estrogen has also been shown to change the total content of cell surface HSPGs and to stimulate their turnover in the mouse uterine epithelium (21). It is widely believed that such changes in epithelial
cell HSPG expression are important for establishment of the
blastocyst-receptive uterine state. The binding interactions of HSPGs
in cell-matrix adhesion may indicate that estrogen-regulated changes in
HSPG expression have an important role in directing endometrial matrix
reorganization and cell spatial arrangements that accompany growth of
the organ. However, little is known about the role of the syndecans in
hormone-regulated uterine growth or about specific mechanisms that
regulate their expression in the uterus.
In this study, we investigated the regulatory effect of estrogen on
syndecan-3 expression in the rat uterus. Analysis of syndecan-3 mRNA and protein indicate that transient changes in syndecan-3 expression occur in response to estrogen treatment. Immunohistochemical analysis of uterine tissue sections showed high levels of expression in
epithelial cells that line the uterine lumen and a change in cell
surface localization of the protein in these cells in response to
estrogen treatment.
Animal Treatments and Tissue Preparations--
All animal
experiments were conducted in accordance with National Institutes of
Health Guidelines for the Care and Use of Laboratory Animals. Female
Harlan Sprague Dawley rats (Buckshire Corp., Perkasie, PA) were
ovariectomized at 21 days of age. Three to five days after surgery,
animals received an intraperitoneal injection of 17
For RNA and protein analysis, animals were killed by decapitation at
defined intervals after injection, and the uteri were stripped of
adhering fat and mesentery, removed, and immediately frozen at the
temperature of liquid nitrogen ( RNA Extraction--
Total RNA was extracted from pooled uterine
samples (approximately three to five uteri/sample) using Tri Reagent
solution (22). Frozen tissue was homogenized in Tri Reagent using a
Polytron homogenizer (Brinkmann Instruments), and the homogenates were phase separated by addition of chloroform and centrifugation at 12,000 × g. RNA was precipitated from the aqueous
phase by addition of 100% isopropanol and pelleted by centrifugation
at 12,000 × g. The final RNA pellet was washed once
with 75% ethanol, briefly dried, and dissolved in FORMazol (Molecular
Research Center). Total RNA was quantified by spectrophotometric
analysis at an absorbance of 260 nm.
Northern Blot--
Total RNA was resolved in a 1%
formaldehyde-agarose gel by electrophoresis at 100 V in a 1×
4-morpholinepropanesulfonic acid buffer (20 mM
4-morpholinepropanesulfonic acid, 8.3 mM NaOAc, 1 mM EDTA, pH 7.0). RNA samples contained ethidium bromide (1 µg) for later visualization under ultraviolet light. An RNA ladder (0.24-9.5 kb; Life Technologies, Inc.) was loaded into one lane of
each gel as molecular weight standard. Gels were photographed under UV
illumination (Eagle Eye System; Stratagene), and the images were used
for densitometric analysis of the 18s rRNA band to correct for loading
inequalities between samples. Resolved RNA was transferred to Magna
0.45-µm nylon membrane (Magna Separation Industries) using a downward
capillary transfer technique (23) in 10× SSC as transfer buffer. RNA
was fixed by baking at 80 °C for 15 min, followed by UV
cross-linking (GS Gene Linker; Bio-Rad).
Northern Blot Hybridization and
Analysis--
32P-Radiolabeled nucleic acid probes were
prepared from a 1-kb EcoRI fragment of cloned rat syndecan-3
cDNA (16). 23 ng of purified cDNA fragment were used for probe
synthesis by random primer labeling (Prime-It II kit; Stratagene) with
Protein Extraction--
Protein was extracted from pooled
samples of frozen uteri (three to six uteri/sample) by homogenization
in a buffer containing 0.1 M Tris (pH 7.5) using a Polytron
homogenizer. Homogenates were centrifuged at 17,000 × g for 30 min at 4 °C, supernatants were saved, and the
pellets were resuspended in buffer containing 0.1 M Tris
(pH 7.5) plus 1% Triton X-100. Pellet extracts were centrifuged as
before, and the supernatants were removed and pooled with the
supernatant from the initial extraction step. Aliquots of mixed
supernatant samples were quantified by spectrophotometric analysis
using a colorimetric dye assay (Bio-Rad).
Immunoblotting--
Extracted protein (25 µg) was denatured at
100 °C in a solution containing 0.5 M Tris-HCl (pH 6.8),
2% SDS, 2% 2-mercaptoethanol, 10% glycerol, and 0.002% bromphenol
blue. Samples were applied to a 6% SDS-polyacrylamide gel and resolved
by electrophoresis (Mini Protean II apparatus; Bio-Rad) at 100 V in a
buffer containing 2.5 mM Tris base (pH 8.3), 192 mM glycine, and 0.1% SDS. After electrophoresis the
proteins were transferred to an Immobilon-P membrane (Millipore) with a
100-V current. Membranes were soaked in a blocking solution of Blotto
plus 1% Tween 20 and then incubated in a 1:250 dilution of polyclonal
rabbit (anti-rat) syndecan-3 antibody (16) in blocking solution at room
temperature using a microhybridization oven. The antibody solution was
removed, and the membrane was washed and then incubated in a 1:3000
dilution of secondary horseradish peroxidase-conjugated goat
(anti-rabbit) IgG (Bio-Rad) in blocking solution. Membranes were again
washed and processed for detection of syndecan-3 protein by incubation in an appropriate substrate solution containing a luminol compound (SuperSignal Substrate; Pierce) for chemiluminescent signal
development. Luminescent membranes were exposed to x-ray film for
signal detection. The intensity of luminescent bands was quantified by
x-ray film densitometry (EagleEye; Stratagene) and used for comparison
of protein content in each sample of uterine protein.
Immunohistochemistry--
Uteri were harvested, and the horns
were separated at the cervix. Each horn was embedded in OCT tissue
medium (Electron Microscopy Services) and frozen. Uterine tissue
sections (8 µm thickness) were generated using a cryostat microtome
(Reichert-Jung Histostat) at Changes in Uterine Syndecan-3 mRNA Levels in Response to
E2--
E2-induced changes in syndecan-3
mRNA content were assessed by Northern blot analysis of RNA samples
extracted from ovariectomized animals over a range of post-hormone
treatment intervals from 0 to 48 h that included both the early
and late uterine growth phases. Hybridization of Northern blots with
radiolabeled rat syndecan-3 probe identified one major 5.9-kb
transcript in the uterus (Fig. 1,
top). Phosphorimage analysis of hybridized blots indicates
that levels of syndecan-3 mRNA increased as early as 2 h and
reached a peak value ~3-fold higher 4 h after E2
administration compared with levels in saline control animals (Fig. 1,
bottom). Transcript levels remained elevated at 18 and
24 h after hormone treatment and then returned to the saline
control level by 48 h. There were no changes detected in
syndecan-3 mRNA content in 24- or 48-h saline-injected control
animals. These data clearly indicate a transient increase in syndecan-3
mRNA content in response to estrogen treatment with peak changes in
mRNA occurring during the early uterine growth phase.
Effect of 17 E2-induced Changes in Uterine Syndecan-3 mRNA
Content Are Transcription but Not Translation Dependent--
Although
the changes in syndecan-3 mRNA are indicative of a direct
hormone-regulated process, it was of interest to determine whether the
increase in mRNA after hormone treatment is the result of a direct
change in transcription of the syndecan-3 gene. To study the
transcription-dependent nature of syndecan-3 mRNA
changes, ovariectomized animals were pretreated with the transcription inhibitor actinomycin D or saline as control before E2
administration 2 h later. Tissues were harvested 4 h after
hormone challenge to determine the effect of the inhibitor at the time
point when mRNA content was estimated to be at the peak level after
estrogen treatment. Hybridization results are shown in Fig.
3. Syndecan-3 mRNA content in animals
pretreated with actinomycin D and challenged with E2,
(Act D + E2) were significantly reduced 2-fold
from transcript levels observed in animals treated with E2
alone (saline + E2). This decrease in mRNA
content by the transcription-inhibitory compound thus indicates that
the syndecan-3 gene is regulated at the level of transcription.
The E2-induced changes in syndecan-3 mRNA,
however, were not similarly affected by pretreatment of animals with
the protein translation inhibitor cycloheximide. Animals were
pretreated with cycloheximide (4 mg/kg) or saline vehicle 2 h
before E2 administration, and mRNA levels were measured
in uterine tissue 4 h after hormone challenge. Syndecan-3 mRNA
content was increased ~2-3-fold in cycloheximide-pretreated animals
(Fig. 3, CHX + E2), which was similar to the
level of change observed in animals pretreated with saline
(saline + E2).
Changes in Uterine Syndecan-3 mRNA Are Steroid Hormone
Specific--
To more specifically investigate hormone responsiveness
of the syndecan-3 gene, ovariectomized animals were treated with
various estrogenic and nonestrogenic steroid hormones, and mRNA
content was determined by Northern blot analysis. Animals were treated with the estrogenic compounds E2, estriol, and tamoxifen or
nonestrogenic steroid compounds, including progesterone,
5 E2-induced Changes in Syndecan-3 mRNA Levels Are
Specific to the Uterus--
Because the observed changes in uterine
syndecan-3 mRNA clearly indicate hormone regulation of gene
transcription, it was of interest to assess whether E2
effected similar changes in other tissues known to contain the estrogen
receptor, including heart (24), liver (25), lung (26), and spleen (27).
All tissues were harvested from the same hormone-treated animals whose
uteri were extracted and used for the initial syndecan-3 Northern blot analysis shown in Fig. 1. The syndecan-3 mRNA transcript was
detected in all tissues analyzed; however, there were no detectable
changes in mRNA levels after E2 administration in any
of these tissues at any postinjection interval (Fig.
5).
Estradiol-induced Temporal and Spatial Changes in Uterine
Syndecan-3 Expression--
To assess changes in expressed syndecan-3
protein during the E2-induced uterine growth response,
Western blot analysis of total uterine protein extracts was
completed. Anti-syndecan-3 antibodies identified a broad high molecular
mass band in the range of 190-250 kDa (Fig.
6). This is consistent with previous observations of the protein in other rat tissues and likely reflects varying degrees of glycanation at potential GAG attachment sites on the
core protein (16). Protein content clearly increased by 8 h and
remained elevated at 20 h after hormone treatment. Levels then
appeared to decline and return to that measured in saline control
animals by 48 h. These changes in syndecan-3 protein content
therefore initiate during the hypertrophic phase but peak early in the
hyperplastic phase of uterine growth and clearly demonstrate transient
regulation of syndecan-3 expression by the estrogenic hormone.
Previous work in the mouse uterus has shown regulated changes in
syndecan-1 expression in vivo during the normal fertility cycle in adult animals (20). To better characterize the cell type-specific expression of syndecan-3 and assess changes in the pattern of uterine syndecan-3 expression, immunohistochemical localization was completed. The same anti-syndecan-3 antibodies were
used to visualize location of the protein in sections of uteri
harvested from saline control and hormone-treated animals. Results show
that syndecan-3 is expressed in epithelial cells of glands in the
endometrial stroma, smooth muscle cells of the myometrium, and
epithelial cells of the perimetrium (Fig.
7). The protein is also highly expressed
in columnar epithelial cells of the endometrium (Fig.
8). Changes in staining intensity in all
cell types indicate that protein levels increase after hormone treatment; however, the most pronounced changes in expression pattern
were observed in the epithelial cells of the endometrium that line the
uterine lumen.
The staining pattern in uteri of saline-treated control animals
indicated a diffuse expression of syndecan-3 along all cell borders
apical to basal (Fig. 8). However, 8 h after E2
treatment, syndecan-3 was diminished significantly at apical cell
borders but was higher in concentration along basolateral membrane
regions. By 20 h, the protein was nearly entirely localized to the
basal cell surfaces of these epithelial cells. At 48 h after
hormone treatment, when the tissue growth response had been completed, the staining profile was again similar to that observed in
saline-treated animals, with syndecan-3 evident on both apical and
basal cell surfaces.
The immature rat uterus has been widely used as a model system to
study the biochemical and molecular mechanisms of steroid hormone
action. Administration of a single physiological dose of E2
(40 µg/kg) induces a biphasic growth response that is rapid in onset,
reaching completion within 48 h (28). The initial phase of the
mitogenic growth involves a series of early hypertrophic responses that
appear 4-6 h after hormone treatment and a series of late hyperplastic
responses that develop 24-30 h after hormone administration (29). The
early responses induced by E2 are characterized by altered
expression of genes encoding for transcription factors (30), oncogenes
(31-33), and growth factors and their receptors (34, 35). Coupled with
the changes in gene expression, dramatic changes in the structure and
organization of the extracellular matrix have also been reported to
occur in the early phases of the growth response (36). Heparan sulfate
proteoglycans are a diverse group of multifunctional integral membrane
proteins that can bind a variety of extracellular ligands, including
matrix adhesion proteins and soluble growth factors. These binding
interactions indicate a potential role for HSPGs in regulated cell
adhesion, morphogenesis, and growth. However, identification of
specific factors that regulate HSPG expression remains unclear. We
report for the first time that syndecan-3 mRNA levels are
controlled by estrogen in the immature rat uterus and that syndecan
proteins are expressed in spatially specific areas in epithelial cells of the endometrium.
Administration of a single physiological dose of E2
increased uterine syndecan-3 mRNA and protein steady-state levels.
Tissue levels of the transcript reach a maximum at 4 h after
hormone administration, a time point that falls within the early phase of uterine growth. Similar analyses of E2 effects on
uterine expression of syndecans-1 and 2 also clearly indicate transient
increases in mRNA and protein content, however, with distinct
temporal patterns.2 These
observations clearly indicate specific E2-regulated
expression of multiple syndecan family members and suggest a role for
these molecules in uterine fertility.
The rapid induction of syndecan-3 mRNA suggests that the syndecan-3
gene is an early growth response gene in the uterus. Analysis of the
dependence of mRNA changes on new gene transcription or protein
translation events supports a direct effect of E2 on
syndecan-3 gene transcription. Pretreatment of animals with actinomycin
D, a transcription inhibitor, significantly blocked the
E2-induced increase in mRNA. However, the translation
inhibitor cycloheximide failed to affect the hormone-induced mRNA
response. The dosages of actinomycin D and cycloheximide used in these
experiments have been shown to be effective in inhibition of
accumulation of other mRNAs and proteins in response to
E2 administration (30, 37) and to block growth-related
uterine morphological changes (36).
Regulated expression of the syndecan-3 gene has been best characterized
during development of the central nervous system in which dramatic
temporal changes in expression correlate with intervals of cell
differentiation (13). Similarly, temporal changes in expression have
been observed during chondrocyte growth intervals in avian limb
formation (38). The molecular factors that regulate proteoglycan gene
expression in these systems have not been elucidated. Although we have
shown in the present studies that syndecan-3 gene transcription is
hormone regulated in the uterus, the specific mechanism by which
E2 elicits this effect is unclear. Complete DNA sequence
analysis of the 5'-flanking region of the rat syndecan-3 gene has not
been completed. Therefore, information on the presence of an estrogen
response element is lacking. Without evidence for specific target sites
for binding of activated steroid hormone receptors, the exact mechanism
of E2 regulation is unknown. It is possible that
E2 induces a change in activity of existing transcription factors, thereby increasing mRNA transcription, or that the hormone significantly affects syndecan-3 mRNA stability, thus elevating mRNA levels through an increase in transcript half-life. Such forms
of control have been described for regulated expression of the
vitellogenin gene in Xenopus hepatocytes after
E2 exposure (39, 40). Effects of actinomycin D on
E2 induction of uterine syndecan-3 mRNA indicate that a
change in transcript stability alone does not account for the observed
mRNA increase and that mRNA changes in the uterus are, at least
in part, transcription dependent.
The present studies show that the syndecan-3 response is hormone
specific. Only steroid ligands that bind the estrogen receptor, including estriol and tamoxifen, induce increased transcript levels in
the uterus. Despite evidence of receptors for nonestrogenic steroids in
the uterus (28, 41, 42), their lack of effect indicates a transcription
response specific to activation of the estrogen receptor. The
glucocorticoid dexamethasone induced a decrease in mRNA content
below that measured in saline control tissues. Dexamethasone has been
shown to down-regulate specific mRNAs in other systems, such as the
PTHrP gene transcript in a human C-cell line (43). Dexamethasone
treatment of S115 mammary carcinoma cells induces a dramatic change in
cell morphology that is accompanied by a loss of syndecan-1 protein
(44). This effect suggests repression of syndecan-1 gene expression by
the glucocorticoid and is consistent with the observed syndecan-3
mRNA down-regulation observed here in the uterus.
In addition to the steroid-specific nature of the uterine syndecan-3
response, there is also tissue specificity. Analysis of other
estrogen-responsive rat tissues clearly indicated the presence of the
major 5.9-kb transcript in heart, liver, lung, and spleen; however,
mRNA levels remained constant 2-48 h after E2
treatment. The lack of a hormone response in these tissues is not
known. This could reflect specific regulatory mechanisms or
differential tissue levels of the estrogen receptor. Although a
complete explanation for differential tissue responsiveness is lacking,
it is clear that the response in the uterus is significant in magnitude
and occurs with a physiological dose of the hormone equivalent to
in vivo circulating levels in cycling adult animals.
Uterine syndecan-3 protein content also increased transiently during
the growth response, with a peak elevation occurring at ~8-20 h
after hormone administration. This temporal change in protein
correlates well with the time course of change in mRNA and further
indicates that regulation of expression is mainly at the level of
transcription. Immunohistochemical staining to visualize syndecan-3
protein in intact uterine sections revealed a high level of expression
in endometrial epithelial cells that line the uterine lumen, with
additional expression detected in glandular epithelial cells of the
endometrial stroma, as well as smooth muscle cells of the myometrium.
Hormone-induced changes in staining intensity in these cells indicate
up-regulation of the protein during the tissue growth response.
However, the most dramatic changes in temporal and spatial expression
occurred in lumenal epithelial cells, where the protein changed from a
more uniform distribution on apical and basolateral cell surfaces in saline-treated tissues to a nearly complete basal localization 20 h after E2 administration. The increase in syndecan-3
levels corresponded to initiation of altered spatial localization in epithelial cells. Similar changes in the histochemical distribution of
syndecan-1 have been described in the mouse uterus during the normal
fertility cycle and early pregnancy (20). Syndecan-1 became localized
to the basal border of endometrial epithelial cells during estrous
cycle intervals when circulating levels of ovarian hormones are
highest. These observations were made in animals with an intact
pituitary-ovary axis; therefore, changes in syndecan-1 expression could
not be attributed solely to estrogen. However, estrogen has been
specifically shown to change the total content of cell surface HSPGs
and to stimulate their turnover in the mouse uterine epithelium (21).
It is widely believed that these hormone-induced changes in epithelial
HSPG expression are important for formation of the blastocyst-receptive
uterine condition. Their high degree of negative charge caused by
sulfated heparan sugars likely promotes binding of important cationic
molecules but may interfere with molecular interactions during
blastocyst implantation (20). Localization of these proteoglycans away from the apical surfaces may facilitate epithelial cell-embryo attachment by decreasing charge-related molecular repulsion.
Basolateral positioning of syndecan-3 might also be crucial for
alterations in epithelial cell interactions with underlying stromal
connective tissue. Limited information is available concerning potential extracellular ligands for syndecan-3. Studies in Schwann cells have indicated direct binding of syndecan-3 to a collagen-like adhesive protein called p200 (45), therefore potentially eliciting a
direct effect on cell adhesion and matrix organization. The presence of
this ligand in the uterus has not been determined. Concentration of
syndecan-3 to basal epithelial borders could also modify actin
cytoskeleton networks either by direct actin binding or through a
coreceptor function in integrin-mediated adhesion. At this time a
direct binding interaction between the cytoplasmic domain of syndecan-3
and the actin cytoskeleton has not been confirmed. Actin microfilament
reorganization might be a consequence of intensified basal surface
expression and might lead to a change in epithelial cell morphology and
polarization. Changes in epithelial cell polarization are evident
during estrogen-induced uterine growth (46) and suggest a potential key
role for syndecan-3 in mediating binding interactions significant for
uterine fertility. Although we have demonstrated estrogen-regulated
changes in syndecan-3 gene transcription and altered protein spatial
expression, determination of binding interactions with cell surface
integrins, matrix adhesion molecules, and cytoskeletal actin in
endometrial epithelial cells will be important to more clearly define
protein function in hormone-regulated uterine fertility.
-estradiol. Administration of a single physiological dose
of 17
-estradiol (40 µg/kg) to ovariectomized immature animals
induced a rapid and transient increase in uterine syndecan-3 mRNA.
Transcript levels reached a peak elevation of 3-fold above saline
control tissues 4 h after hormone administration. Inhibition of
message up-regulation by actinomycin D but not cycloheximide indicated a hormone response dependent on RNA transcription but not new protein
synthesis. The estrogenic ligands estriol and tamoxifen were also
effective at raising syndecan-3 mRNA levels; however, nonestrogenic
ligands, including progesterone, 5
-dihydrotestosterone, and
dexamethasone, failed to stimulate a change in mRNA levels. Hormone-induced changes in mRNA led to transient changes in
syndecan-3 protein content and significant alteration in the temporal
and spatial expression in endometrial epithelial cells. Collectively, these data show that the steroid hormone 17
-estradiol,
regulates transcription of the syndecan-3 gene in the uterus via an
estrogen receptor-dependent mechanism. This
estrogen-regulated expression of syndecan-3 may play an important role
in changes in tissue ultrastructure crucial for proper uterine growth.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-estradiol
(E2; 40 µg/kg in a 0.9% NaCl, 0.4% ethanol vehicle).
Control animals received vehicle only. For experiments in which animals
were treated with gene transcription or protein translation inhibitors,
intraperitoneal injection of actinomycin D (8 mg/kg) or cycloheximide
(4 mg/kg) was administered 2 h before E2 or control
vehicle injection. Additional treatment compounds used for steroid
specificity analysis were administered at the following doses:
dexamethasone (600 µg/kg), estriol (40 µg/kg), tamoxifen (20 mg/kg), progesterone (4 mg/kg), and 5
-dihydrotestosterone (400 µg/kg). All chemicals were obtained from Sigma.
270 °C). For tissue specificity
experiments, additional tissues were removed from these same animals
and handled similarly.
[32P]dCTP (3000 Ci/mmol; Amersham Pharmacia Biotech)
as isotope. Northern blots were prehybridized at 65 °C in a solution
containing 5× saline/sodium phosphate/EDTA, 50% formamide, 0.5% SDS,
100 µg/ml denatured salmon sperm DNA, and 5× Denhardt's reagent.
Prehybridized blots were incubated at 65 °C in a hybridization
solution (5× saline/sodium phosphate/EDTA, 0.5% SDS, 100 µg/ml
denatured salmon sperm DNA, 5× Denhardt's reagent, 10% dextran
sulfate) and the 32P-labeled probe (~1.2-2.1 × 109 dpm/µg DNA). Membranes were washed under standard
conditions and exposed to x-ray film (Fuji RX). For quantitative
comparisons of mRNA levels, hybridization signals were quantified
by direct measurement of membrane-bound radioactivity by phosphorimage
analysis (Molecular Dynamics). Nonspecific background counts per minute in each lane were subtracted from the total counts per minute for
radiolabeled mRNA bands to yield net counts per minute. Net counts
were used for quantification of mRNA levels and direct comparison
between RNA samples.
25 °C and placed on latex-coated
slides (Cell Point Scientific). Tissue sections were fixed in 3%
paraformaldehyde in phosphate-buffered saline, rinsed in
phosphate-buffered saline, and then incubated with a 1:50 dilution of
polyclonal rabbit (anti-rat) syndecan-3 antibodies in blocking solution
(0.2 M NaCl, 0.1 M Tris, pH 7.7, 5% nonfat
milk) at room temperature. The antibody solution was removed, and
sections were washed with blocking agent and then incubated in a 1:100
dilution of Texas Red-conjugated goat (anti-rabbit) IgG (Molecular
Probes) for a minimum of 1 h at room temperature. Tissue sections
were washed as before and then rinsed briefly with 0.1 M
phosphate-buffered saline. Sections were examined on an inverted
microscope equipped for epifluorescence (Axiovert 35; Zeiss) using the
appropriate filters for excitation of the Texas Red fluorescent marker
and visualization of emitted light. Sections were photographed, and the
images were used for identification of cell type expression of
syndecan-3 and analysis of differences in expression pattern between
uterine tissue samples. As a control, uterine sections from the same
animals were incubated in blocking agent alone and then exposed to
Texas Red-conjugated secondary antibodies. Staining in control sections
indicated a weak, diffuse background fluorescence that was similar
between saline- and hormone-treated animals.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (45K):
[in a new window]
Fig. 1.
Time course of change in uterine syndecan-3
mRNA. Top, total RNA (12 µg/lane) from uteri of
ovariectomized animals 0-48 h after treatment with saline (control) or
E2 (40 µg/kg) was separated by formaldehyde-agarose gel
electrophoresis and blotted onto a nylon membrane. Blots were
hybridized with 32P-labeled probe for detection of the
5.9-kb syndecan-3 transcript. Data shown are typical of three analyses
using total RNA isolated from different animals. Bottom,
hybridization signals from the blot depicted above were quantitated by
phosphorimage analysis. Counts per minute represented for each sample
were corrected for background radioactivity and normalized as described
under "Experimental Procedures."
-Estradiol on Syndecan-3 mRNA Is Dose
Dependent--
To determine whether the effect of E2 on
syndecan-3 mRNA is dose dependent and thus indicative of a true
hormone-regulated process, ovariectomized animals were treated with
increasing doses of E2, and levels of syndecan-3 mRNA
were analyzed by Northern blot. Because the initial Northern blot
analysis (Fig. 1) indicated that the induced change in syndecan-3
mRNA content was greatest 4 h after hormone treatment, all
uterine tissues were extracted at this time point after either
E2 or saline control injections. Little or no change in
syndecan-3 mRNA was detected at E2 dosages of <0.4
µg/kg (Fig. 2, top).
However, mRNA levels increased to 43% of the maximum hormone
response at a dose of 0.4 µg/kg (Fig. 2, bottom). Doses of
4 and 40 µg/kg produced equivalent increases in mRNA, indicating
a maximum response level had been reached. The characteristic
dose-response curve from the analysis (Fig. 2, bottom) is
consistent with a response that is controlled by physiological levels
of estradiol.
View larger version (39K):
[in a new window]
Fig. 2.
Dose-dependent changes in uterine
syndecan-3 mRNA content. Top, total RNA (12 µg/lane) isolated from ovariectomized rats 4 h after treatment
with either saline (control) or E2 at the indicated dosages
was separated by formaldehyde-agarose gel electrophoresis and blotted
onto a nylon membrane. Blots were hybridized to detect the 5.9-kb
syndecan-3 transcript. Bottom, hybridization signals from
the blot depicted above were quantitated by phosphorimage analysis.
Results are presented as percentage of the maximum change induced by
E2 at the 40-µg/kg dose.
View larger version (50K):
[in a new window]
Fig. 3.
Effects of actinomycin D and cycloheximide on
estrogen-induced changes in uterine syndecan-3 mRNA levels.
Top, total RNA was isolated from ovariectomized animals
under the following treatment protocols: saline pretreatment
(Saline) followed by saline (S) or E2
challenge (E2) 2 h later, actinomycin D pretreatment
(Act D) followed by saline or E2 challenge
2 h later, and cycloheximide (CHX) pretreatment
followed by saline or E2 challenge 2 h later. All
tissues were harvested 4 h after the saline or E2
challenge. 12 µg of RNA were separated by gel electrophoresis and
analyzed by Northern blot. Hybridization of radiolabeled probe to the
5.9-kb transcript is represented. Results are typical of more than
three blot analyses with RNA from different animals in two separate
experiments. Bottom, quantitated hybridization signals from
the blot shown at top.
-dihydrotestosterone, and dexamethasone at the indicated effective
dosages (Fig. 4). Uterine tissues were
harvested from all treatment groups 4 h after hormone injections
and processed for Northern blot analysis. Of the steroid hormones
administered, only the estrogenic compounds E2, estriol,
and tamoxifen induced increases in syndecan-3 mRNA content that
were ~2-3-fold above levels measured in saline control animals for
all three compounds (Fig. 4, bottom). The nonestrogenic sex
steroid compounds progesterone and 5
-dihydrotestosterone caused no
change in syndecan-3 mRNA over the same interval. However, treatment with the glucocorticoid agonist dexamethasone produced a
decrease in mRNA levels, an ~80% reduction, compared with levels in tissues from saline control animals.
View larger version (40K):
[in a new window]
Fig. 4.
Steroid specificity of changes in uterine
syndecan-3 mRNA. Top, total RNA (12 µg/lane) was
isolated from ovariectomized animals 4 h after treatment with
saline, estradiol (E2, 40 µg/kg), estriol
(E3, 40 µg/kg), tamoxifen (TAM, 20 mg/kg), progesterone (PRG, 4 mg/kg),
5 -dihydrotestosterone (DHT, 400 µg/kg), or
dexamethasone (DEX, 600 µg/kg) and analyzed by Northern
blot to detect the 5.9-kb syndecan-3 transcript. Bottom,
quantitated hybridization signals from the blot depicted above.
View larger version (77K):
[in a new window]
Fig. 5.
Tissue specificity of estrogen-induced
changes in syndecan-3 mRNA. Total RNA (12 µg/lane) was
isolated from the uteri, heart, liver, lung, and spleen of
ovariectomized animals 0-48 h after treatment with saline (control) or
E2 (40 µg/kg) and analyzed by Northern blot to detect the
5.9-kb syndecan-3 transcript.
View larger version (78K):
[in a new window]
Fig. 6.
Time course of change in uterine syndecan-3
protein. Protein was extracted from the uteri of ovariectomized
animals at the indicated intervals 0-48 h after treatment with either
saline (control) or E2 (40 µg/kg). Protein samples (25 µg/lane) were size separated by SDS-polyacrylamide gel
electrophoresis and transferred to an Immobilon-P membrane via Western
blot. Binding of anti-syndecan-3 antibodies was detected through
chemiluminescence as described under "Experimental Procedures."
Analysis indicated the presence of a high molecular mass band of
~190-250 kDa. Similar analyses were repeated more than five times
with protein extracts from different animals and yielded similar
results.
View larger version (118K):
[in a new window]
Fig. 7.
Histochemical analysis of syndecan-3 protein
in the uterus. Staining of 8-µm cryosection of rat uterine
tissue for visualization of the syndecan-3 protein using
anti-syndecan-3 antibodies indicated the presence of the protein in
epithelial cells of the perimetrium (P), smooth muscle cells
of the myometrium (M), glandular epithelial cells
(g) of the stroma (S), and blood vessel
endothelium (b). Scale bar, 20 µm.
View larger version (147K):
[in a new window]
Fig. 8.
Histochemical analysis of temporal and
spatial changes in syndecan-3 protein expression. Uteri were
removed from saline-treated (control) and E2-treated (40 µg/kg) ovariectomized animals, and 8-µm frozen sections were
prepared for immunodetection of syndecan-3 protein. Binding of
anti-syndecan-3 antibodies was detected indirectly using Texas
Red-conjugated mouse (anti-rabbit) IgG. Staining of sections shows the
expression pattern specifically in the endometrial epithelium
(E); the uterine lumen (L) and endometrial stroma
(S) are also indicated. Scale bar, 20 µm.
Panels reflect a time course of hormone response intervals:
1, saline-treated control animals; staining is evident on
apical (a) and basal (b) cell borders;
2, 2 h after E2 treatment; 3,
6 h after E2 treatment; 4, 8 h after
E2 treatment; staining at the apical cell surface
(a) is diminished and more intense at the basolateral
surfaces (b); 5, 12 h after E2
treatment; almost exclusive basal staining is evident (b);
6, 20 h after E2 treatment; basal
localization still evident; 7, 48 h after
E2 treatment; return to a staining pattern similar to that
in saline tissues with reappearance of protein at apical cell borders
(a); 8, control immunofluorescent tissue
preparation; 12-h post-E2 uterus exposed only to
fluorescent secondary antibodies.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ To whom correspondence should be addressed: Dept. of Biology, Villanova University, Mendel Hall, 800 Lancaster Ave., Villanova, PA 19085. Tel.: 610-519-4869; Fax: 610-519-7863; E-mail: louise.russo@villanova.edu.
Published, JBC Papers in Press, October 6, 2000, DOI 10.1074/jbc.M004106200
2 L. A. Russo, S. P. Calabro, T. A. Filler, D. J. Carey, and R. M. Gardner, unpublished observations.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
HSPG, heparan
sulfate proteoglycan;
GAG, glycosaminoglycan;
E2, 17-estradiol;
kb, kilobase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Murphy, C. R., and Rogers, A. W. (1981) Cell. Biophys. 3, 305-320[Medline] [Order article via Infotrieve] |
2. | Bernfield, M., Kokenyesi, R., Kato, M., Hinkes, M. T., Spring, J., Gallo, R. L., and Loose, E. J. (1992) Annu. Rev. Cell Biol. 8, 365-393[CrossRef] |
3. | Carey, D. A. (1997) Biochem. J. 327, 1-16[Medline] [Order article via Infotrieve] |
4. | Klagsbrun, M. (1990) Curr. Opin. Cell Biol. 2, 857-863[Medline] [Order article via Infotrieve] |
5. | Rapraeger, A., Krufka, A., and Olwin, B. (1991) Science 252, 1705-1708[Medline] [Order article via Infotrieve] |
6. |
Gitay-Goren, H.,
Soker, S.,
and Vlodavsky, I.
(1992)
J. Biol. Chem.
267,
6093-6098 |
7. | Saunders, S., and Bernfield, M. (1988) J. Cell Biol. 106, 423-430[Abstract] |
8. |
Barkalow, F. J.,
and Schwarzbauer, J. E.
(1991)
J. Biol. Chem.
266,
7812-7818 |
9. |
Kouzi-Koliakos, K.,
Koliakos, G. G.,
Tsilibary, E. C.,
Furcht, L. T.,
and Charonis, A. S.
(1989)
J. Biol. Chem.
264,
17971-17978 |
10. | LeBaron, R. G., Esko, J. D., Woods, A., Johansson, S., and Hook, M (1988) J. Cell Biol. 106, 945-952[Abstract] |
11. | Woods, A., and Couchman, J. R. (1994) Mol. Biol. Cell 5, 183-192[Abstract] |
12. | Baciu, P. C., and Goetinck, P. F. (1995) Mol. Biol. Cell 6, 1503-1513[Abstract] |
13. |
Carey, D. J.,
Conner, K.,
Asundi, V. K.,
O'Mahony, D. J.,
Stahl, R. C.,
Showalter, L. J.,
Cizmeci-Smith, G.,
Hartman, J.,
and Rothblum, L. I.
(1997)
J. Biol. Chem.
272,
2873-2879 |
14. | Reyes, A. A., Akeson, R., Brezina, L., and Cole, G. J. (1990) Cell Regul. 1, 567-576[Medline] [Order article via Infotrieve] |
15. | Couchman, J. R., and Woods, A. (1996) J. Cell. Biochem. 61, 578-584[CrossRef][Medline] [Order article via Infotrieve] |
16. | Carey, D. J., Evans, D. M., Stahl, R. C., Asundi, V. K., Conner, K. J., Garbes, P., and Cizmeci-Smith, G. (1992) J. Cell Biol. 117, 191-201[Abstract] |
17. |
Elenius, K.,
Maata, A.,
Salmivirta, M.,
and Jalkanen, M.
(1992)
J. Biol. Chem.
267,
6435-6441 |
18. |
Cizmeci-Smith, G.,
Stahl, R. C.,
Showalter, L. J.,
and Carey, D. J.
(1993)
J. Biol. Chem.
268,
18740-18747 |
19. |
Cizmeci-Smith, G.,
Langan, E.,
Youkey, J.,
Showalter, L. J.,
and Carey, D. J.
(1997)
Arterioscler. Thromb. Vasc. Biol.
17,
172-180 |
20. | Potter, S. W., and Morris, J. E. (1992) Anat. Rec. 234, 383-390[Medline] [Order article via Infotrieve] |
21. |
Morris, J. E.,
Potter, S. W.,
and Gaza-Bulseco, G.
(1988)
J. Biol. Chem.
263,
4712-4718 |
22. | Chomczynski, P. (1993) BioTechniques 15, 53-57 |
23. | Chomczynski, P. (1992) Anal. Biochem. 201, 134-139[Medline] [Order article via Infotrieve] |
24. | Klangkalya, B., and Chan, A. (1988) Life Sci. 42, 2307-2314[CrossRef][Medline] [Order article via Infotrieve] |
25. | Freychuss, B. L., Sahlin, L., Masironi, B., and Reiksson, H. (1994) J. Endocrinol. 142, 285-298[Abstract] |
26. | Thuresson-Klein, A., Moawad, A. H., and Hedqvist, P. (1985) Am. J. Obstet. Gynecol. 151, 506-514[Medline] [Order article via Infotrieve] |
27. | Myers, M. J., Heim, M. C., Hirsch, K. S., Queener, S. F., and Petersen, B. H. (1986) Life Sci. 39, 313-320[Medline] [Order article via Infotrieve] |
28. | Anderson, J. N., Clark, J. H., and Peck, E. J. (1972) Biochem. Biophys. Res. Commun. 48, 1460-1468[Medline] [Order article via Infotrieve] |
29. | Kaye, A. M., Sheratzky, D., and Lindner, H. R. (1972) Biochim. Biophys. Acta 261, 475-486 |
30. | Suva, L. J., Harm, S. C., Gardner, R. M., and Thiede, M. A. (1991) Mol. Endocrinol. 5, 829-835[Abstract] |
31. | Webb, D. K., Moulton, B. C., and Khan, S. A. (1990) Biochem. Biophys. Res. Commun. 168, 721-726[Medline] [Order article via Infotrieve] |
32. | Loose-Mitchell, D. S., Chiappetta, C., and Stancel, G. M. (1988) Mol. Endocrinol. 2, 946-951[Abstract] |
33. | Murphy, L. M., Murphy, L. C., and Friesen, H. G. (1987) Endocrinology 120, 1882-1888[Abstract] |
34. | DiAugustine, R. P., Petrusz, P., Bell, G. I., Brown, C. F., Korach, K. S., McLachlan, J. A., and Teng, C. T. (1988) Endocrinology 122, 2355-2363[Abstract] |
35. |
Mukku, V. R.,
and Stancel, G. M.
(1985)
J. Biol. Chem.
260,
9820-9824 |
36. | Pastore, G. N., DiCola, L. P., Dollahon, N. R., and Gardner, R. (1992) Biol. Reprod. 47, 83-91[Abstract] |
37. | Thiede, M. A., Harm, S. C., Hasson, D. M., and Gardner, R. M. (1991) Endocrinology 128, 2317-2323[Abstract] |
38. | Gould, S. E., Upholt, W. B., and Kosher, R. A. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 3271-3275[Abstract] |
39. | Baker, H. J., and Shapiro, D. J. (1978) J. Biol. Chem. 253, 4521-4524[Abstract] |
40. | Brock, M. L., and Shapiro, D. J. (1983) Cell 34, 207-214[Medline] [Order article via Infotrieve] |
41. | Stack, G., and Gorski, J. (1985) Endocrinology 117, 2024-2032[Abstract] |
42. | Dauvois, S., and Parker, M. G. (1993) in Steroid Hormone Action (Parker, M. G., ed) , pp. 166-185, Oxford University Press, New York |
43. |
Ikeda, C. L.,
Weir, E. C.,
Mangin, M.,
and Broadus, A. E.
(1989)
J. Biol. Chem.
264,
15743-15746 |
44. | Leppa, S., Mali, M., Miettinen, H., and Jalkanen, M. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 93-96 |
45. |
Chernousov, M. A.,
Stahl, R. C.,
and Carey, D. J.
(1996)
J. Biol. Chem.
271,
13844-13853 |
46. | Schlafke, S., Welsh, A. O., and Enders, A. C. (1985) Anat. Rec. 212, 47-56[Medline] [Order article via Infotrieve] |