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Molecular Endocrinology, doi:10.1210/me.2003-0014
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Molecular Endocrinology 18 (1): 214-227
Copyright © 2004 by The Endocrine Society

Glucocorticoids Control ß-Catenin Protein Expression and Localization through Distinct Pathways that Can Be Uncoupled by Disruption of Signaling Events Required for Tight Junction Formation in Rat Mammary Epithelial Tumor Cells

Yi Guan, Nicola M. Rubenstein, Kim L. Failor, Paul L. Woo and Gary L. Firestone

Department of Molecular and Cell Biology and The Cancer Research Laboratory, University of California at Berkeley, Berkeley, California 94720-3200

Address all correspondence and requests for reprints to: Gary L. Firestone, Department of Molecular and Cell Biology, 591 LSA, University of California at Berkeley, Berkeley, California 94720-3200. E-mail: glfire{at}uclink4.berkeley.edu.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
In Con8 rat mammary epithelial tumor cells, the synthetic glucocorticoid dexamethasone stimulates the remodeling of tight junctions and adherens junctions before formation of highly sealed tight junctions. In this study, the expression and localization of key components of the apical junction were examined as potential targets of glucocorticoid signaling. Western blot and RT-PCR demonstrated that dexamethasone up-regulated ß-catenin protein and transcript expression and nearly ablated ß-catenin phosphorylation under conditions that led to a significant increase in monolayer transepithelial resistance. Indirect immunofluorescence revealed that dexamethasone treatment also caused ß-catenin to localize predominantly at the cell membrane rather than the nucleus. The glucocorticoid regulation of ß-catenin expression and localization was not a consequence of dexamethasone inhibition of cell growth, because both responses were unaltered in the presence of hydroxyurea. The steroid induction of ß-catenin expression and localization can be uncoupled by altering the function of signaling pathways needed for tight junction formation. Expression of dominant-negative RasN17 abolished dexamethasone up-regulation of ß-catenin protein expression without affecting its localization at the membrane. In contrast, exogenous treatment or constitutive production of TGF{alpha} abolished the dexamethasone-induced alteration of ß-catenin localization without affecting the dexamethasone stimulation of ß-catenin expression. Taken together, our results demonstrate that glucocorticoids control ß-catenin at two distinct levels of cellular regulation that differ in their cell signaling requirements for the glucocorticoid regulation of mammary epithelial junctional dynamics.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
INTERCELLULAR ADHESION IS crucial for the assembly and maintenance of the three-dimensional architecture of tissues (1, 2, 3). Cells interact with each other and/or the extracellular matrix through adhesive molecules such as cadherin, Ig, selectin, and integrin, as well as through the formation of specific protein complex structures such as adherens junctions, tight junctions, desmosomal junctions, and gap junctions (1). The adherens junction is localized laterally at the apical portion of epithelial cells, which can experience contractile or mechanical forces. E-cadherin is the transmembrane component of adherens junctions, and the extracellular regions of E-cadherin that contact neighboring cells form antiparallel, calcium-dependent structures and directly mediate cell-cell adhesion (4). The cytoplasmic domain of E-cadherin binds to ß-catenin or plakoglobin. ß-Catenin binds to {alpha}-catenin, which in turn connects the complex to the cytoskeleton (4, 5, 6). {alpha}-Catenin interacts with other actin-binding proteins, specifically, {alpha}-actinin, vinculin, and the known intracellular tight junction proteins zonula occludens 1 and 2 (ZO-1 and ZO-2) (7, 8, 9, 10, 11, 12, 13). Also on the lateral membrane, immediately apical to the adherens junction, lies the tight junction (for review see Ref. 14). The tight junction regulates paracellular permeability by forming a regulated barrier between cells (15) and defines cell polarity by acting as a boundary within the plasma membrane, separating apical and basolateral membrane proteins (16, 17, 18). The known protein components of the tight junction include transmembrane proteins such as occludin (19), the claudin family (20, 21, 22), and junctional adhesion molecule (23), as well as cytoplasmic proteins such as ZO-1 (12), ZO-2 (13), ZO-3 (24), cingulin (25), 7H6 antigen (26), symplekin (27), and rab13 (28).

It has been established that several of the proteins involved in adherens and tight junction formation can function in other signaling pathways when not recruited to sites of cell-cell contact. For example, ß-catenin has been shown to have important roles in mediating cellular adhesion and in activating transcription (29, 30). ß-Catenin is homologous to the armadillo protein of Drosophila and is a downstream effector of the Wingless/Wnt signaling pathway, which can be activated by progesterone in the mammary gland ((31, 32, 33); http://www.stanford.edu/~rnusse/wntwindow.html). Activation of the Wnt signaling pathway inhibits glycogen synthase kinase 3 (GSK3) activity, which in turn reduces ß-catenin phosphorylation. One cellular consequence of decreased phosphorylation is the stabilization of ß-catenin protein in the cytoplasm and its subsequent nuclear import to regulate transcription (34, 35). Nuclear ß-catenin binds to members of the T cell factor/lymphoid enhancing factor family of transcription factors and stimulates gene expression (36, 37, 38, 39, 40, 41, 42).

Appropriate ß-catenin localization at the adherens junction as well as regulation of ß-catenin stability are crucial for the maintenance of normal epithelial cells. The regulation of ß-catenin is controlled at various levels, such as differential expression of cadherins (43, 44, 45), wnt gene expression (46), or axin/ß-catenin/PP2A /adenomatous polyposis coli complex formation (47, 48, 49, 50, 51, 52, 53, 54). Unregulated ß-catenin expression, or expression of mutant forms of this protein, can lead to cell transformation in vitro and in vivo (55, 56, 57).

Mammary epithelial cells are a useful model system with which to investigate the role of ß-catenin signaling in the context of hormone-controlled mechanisms. The development, differentiation, and proliferation of mammary cells are stringently regulated by systemic and paracrine secreted hormones, locally acting growth factors, and extracellular matrix components (58, 59, 60, 61, 62). Conceivably, the same sets of extracellular cues also control aspects of mammary cell adhesion, because remodeling of the epithelial cell adhesion complex is a general feature underlying the hormone-mediated development of the mammary gland. In the early developmental stages, an increase of expression of desmosomal and tight junction molecules are associated with the ductal mammary epithelial elongation and cavity formation (63, 64). At onset and establishment of lactation, which is progesterone and glucocorticoid dependent (65), an increase in tight junction organization and a decrease in permeability of the mammary epithelium are critical to prevent paracellular leakage of milk components from the alveolar lumen into the mammary gland stroma (66, 67, 68). During mammary gland involution at the end of lactation, one of the first observable changes is the loss of intercellular adhesion (69, 70). Furthermore, disruption of cell adhesion is also reported in pathological conditions such as mastitis (71, 72) and mammary gland tumorigenesis (73, 74, 75). These studies demonstrate that the precise regulation of cell-cell interactions is an integral aspect of maintaining the normal structure and function of the mammary gland.

Using the Con8 rat mammary tumor epithelial cell line derived from a 7,12-dimethylbenz(a)anthracence-induced rat mammary adenocarcinoma (76, 77), we demonstrated that glucocorticoids recruit tight junction and adherens junction components to areas of cell-cell contact and stimulate tight junction formation (78, 79). In this study, we examined the effects of dexamethasone treatment on adherens junction proteins and show, for the first time, that glucocorticoids not only change ß-catenin protein levels, but also alter its phosphorylation and subcellular localization. Furthermore, we show that expression and localization of ß-catenin are part of different signaling pathways that are required for the remodeling of the apical junction and formation of tight junctions. We propose that the multilevel regulation of ß-catenin by glucocorticoids is an important step leading toward formation of both adherens and tight junctions.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Dexamethasone Up-Regulates ß-Catenin Protein Level in Rat Mammary Epithelial Tumor Cells
Our previous results showed that dexamethasone, a synthetic glucocorticoid, induced the formation of tight junction and adherens junction in Con8 rat mammary epithelial tumor cells (78). To detect potential dexamethasone-regulated target(s) at the adherens junction, Con8 cells were treated with or without 1 µM dexamethasone for 5 d, and the expression levels of several adherens junction proteins were examined by Western blots. Over the same time course, the transepithelial electrical resistance (TER) was measured to monitor tight junction sealing. Consistent with our previous results, dexamethasone induced a significant increase in TER over the 5-d time course (Fig. 1AGo). Concurrent with dexamethasone-induced TER increase, ß-catenin protein levels were also up-regulated. The effect of dexamethasone was specific because the level of E-cadherin and {alpha}-catenin, two other adherens junction proteins, remained unchanged (Fig. 1BGo). Approximately equal loading was confirmed by the similar amount of actin detected in each sample. ß-Catenin protein levels obtained from three independent experiments in dexamethasone-treated and untreated cells were quantified and the results depicted in Fig. 1CGo. The 2- to 3-fold increase in ß-catenin protein in steroid-treated cells was statistically significant at all three time points examined (P < 0.05).



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  		Fig. 1. Dexamethasone (Dex) Mediates Up-Regulation of ß-Catenin Protein Expression and Stimulation of TER in Con8 Mammary Epithelial Tumor Cells

Confluent monolayers of Con8 cells were grown on filter supports and treated with or without 1 µM dexamethasone for 5 d. A, The TERs were measured and the ohm·cm2 calculated, as described in Materials and Methods. The results were averages of three separate experiments. B, Cell lysates were normalized for total protein, electrophoretically fractionated, and transferred onto nitrocellulose membranes. The membranes were probed for ß-catenin, E-cadherin, {alpha}-catenin, or actin protein expression. C, Data from three independent Western blot experiments were quantified with the Scion imaging program and analyzed for statistical significance. Asterisks indicate significance at P < 0.05.

 
Dexamethasone up-regulated ß-catenin protein levels in a dosage-dependent way (data not shown). The half-maximal concentration of dexamethasone that up-regulated ß-catenin protein levels was approximately 1 nM, which is close to the half-maximal binding of this steroid to Con8 cell glucocorticoid receptors (76). Moreover, treatment with either 5 µM or 10 µM RU486, a glucocorticoid receptor antagonist, abolished dexamethasone up-regulation of ß-catenin protein expression (Fig. 2Go). These results indicate that the dexamethasone up-regulation of ß-catenin protein is a glucocorticoid receptor-mediated process.



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  		Fig. 2. The Glucocorticoid Receptor Antagonist RU486 Prevents Dexamethasone (Dex) Stimulation of ß-Catenin Protein

Serum-starved Con8 cells were grown for 3 d with or without 1 µM dexamethasone in the presence or absence of either 5 µM or 10 µM RU486. Production of ß-catenin protein in total cell lysates was analyzed by Western blot as described in Fig. 1Go. The level of tubulin protein was used as a gel loading control.

 
Given the transcriptional mechanism of glucocorticoid receptor action (80, 81), we determined whether dexamethasone stimulates ß-catenin transcript levels. Con8 cells were treated with or without 1 µM dexamethasone for 24 or 48 h, total RNA was isolated, and ß-catenin transcript levels were detected by RT-PCR using the primers described (see Materials and Methods). As shown in Fig. 3AGo, dexamethasone stimulated the level of ß-catenin transcripts within 24 h of steroid treatment. Approximately equal amounts of RNA inputs were confirmed by the relatively constant amounts of amplified glyceraldehydes-3-phosphate dehydrogenase (GAPDH) (Fig. 3AGo). By 48 h in glucocorticoids, there is nearly a 4-fold increase in ß-catenin transcripts compared with cells cultured for the same time period without added steroid (Fig. 3Go, B and C).



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  		Fig. 3. Dexamethasone (Dex) Up-Regulates ß-Catenin Transcripts

A, Con8 cells were grown on plastic supports and treated with or without 1 µM dexamethasone for 24 h. Total RNA was isolated from each sample and cDNA was generated by reverse transcription. The transcript levels of ß-catenin and GAPDH were amplified by PCR reaction, and the generated cDNA bands were visualized with a 1% agarose gel. B, After 48 h of dexamethasone treatment, RNA was isolated, and RT-PCR was performed with oligonucleotide primers specific for ß-catenin and 18S rRNA transcripts. The amplified cDNA was visualized by silver staining. C, Three independent RT-PCR experiments examining ß-catenin transcript levels at the 48-h time point were quantified using the Scion imaging program and analyzed for statistical significance. The statistically significant (P = 0.042) results are indicated by an asterisk.

 
Dexamethasone Alters the Distribution of ß-Catenin Protein in Rat Mammary Epithelial Tumor Cells
ß-Catenin is not only a major component of adherens junction at the cell membrane but can also function as a transcriptional regulator in the nucleus (41, 82, 83, 84). The localization of ß-catenin is tightly controlled and determines the major role of this molecule upon various extracellular stimuli (85, 86, 87, 88, 89). To test whether glucocorticoids control ß-catenin protein localization, in addition to its effect on expression, confluent Con8 cells were treated with or without 1 µM dexamethasone for 5 d, and the subcellular distribution of ß-catenin was characterized by indirect immunofluoresence staining and confocal microscopy. As a positive control for the effects of glucocorticoids, the localization of the ZO-1 tight junction protein was also examined in a parallel set of cell cultures. In the absence of dexamethasone, ß-catenin localized predominantly to the nucleus, and as we described previously (78, 79), ZO-1 staining was spotty and discontinuous (Fig. 4AGo; -DEX). Upon 24 h of dexamethasone treatment, ß-catenin and ZO-1 proteins began to appear in a continuous manner on the cell membrane. At d 5 in steroid, ß-catenin and ZO-1 proteins were localized to the cell junctions and displayed a cobblestone pattern (Fig. 4AGo; +DEX). While there were increased amounts of ß-catenin localized to the cell periphery, the nuclear staining of ß-catenin was greatly reduced. In contrast, in untreated cells ß-catenin remained mostly in the nucleus. Nuclear localization of ß-catenin staining was confirmed by 4,6-diamidino-2-phenylindole (DAPI) stain and control secondary antibody staining (Fig. 4BGo).



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  		Fig. 4. Dexamethasone (Dex) Treatment Regulates ß-Catenin’s Subcellular Localization

A, Con8 cells were grown on filter supports and confluent cells were treated with or without 1 µM dexamethasone. At the indicated times, cells were fixed and the localization of ß-catenin and ZO-1 proteins were analyzed by immunofluorescence microscopy. B, DAPI staining was used to assess the nuclear localization of ß-catenin in 24-h glucocorticoid-treated cells. The left panels include anti-ß-catenin antibodies in the immunofluorescence analysis, and the right panels are a control that employ only secondary antibodies. C, Cells were treated with or without 1 µM dexamethasone for 5 d, and Z-planes were analyzed by confocal microscopy using a Zeiss Confocal Microscope, and LSM analyzing software. D, Cells were grown on tissue culture plates and treated for 5 d in the presence or absence of 1 µM dexamethasone. Cells were then biochemically fractionated, and each fraction was normalized to the treatment conditions. Complete lysate and fractions were separated by SDS-PAGE and probed with antibodies for ß-catenin or actin.

 
ß-Catenin localization was also analyzed in dexamethasone-treated and untreated cells by confocal microscopy. Cells were grown on filters and treated with or without steroid for 5 d and then stained for actin and ß-catenin. Analysis of the xz-plane of actin staining revealed that in the absence of dexamethasone treatment, the cells formed disorganized multilayers (Fig. 4CGo, -Dex). ß-Catenin staining appeared mostly as circular areas that overlapped with the nuclear DAPI stain (DAPI staining not shown). In contrast, after 5 d of dexamethasone treatment, the cells formed a monolayer, and the actin was highly organized into perijunctional F-actin rings. In the xz-plane, these F-actin rings resemble vertical lines (Fig. 4CGo, +Dex, actin). Under these conditions, ß-catenin also localized to the lateral membranes, thus also appearing as vertical lines in the micrograph (Fig. 4CGo, +Dex, ß-catenin). This experiment can not distinguish whether ß-catenin is redistributed from the nucleus to the membrane, or whether newly produced ß-catenin is maintained in the cytoplasm and recruited to cell-cell contacts.

To confirm the immunofluorescence results, dexamethasone-treated and untreated cells were biochemically fractionated into total membranes, cytoplasm, and nuclei by differential ultracentrifugation as described in Materials and Methods. As expected, ß-catenin protein increased in whole-cell lysates from steroid-treated cells compared with whole-cell lysates from untreated cells (Fig. 4DGo, Input, -/+Dex). Consistent with the immunofluorescence results, in dexamethasone-treated cells, ß-catenin was enriched in the membrane fraction and depleted in the nuclear fraction, compared with untreated control fractions (Fig. 4DGo, Mem -/+ Dex). In each biochemical fraction, total protein samples were normalized to the level of actin (Fig. 4DGo, lower panel).

Dexamethasone Up-Regulation of ß-Catenin Protein Level and Alteration of ß-Catenin Localization Are Independent of Growth Inhibition
We previously reported that glucocorticoids induce a G1 cell cycle arrest of Con8 cells (90). To test whether the dexamethasone effects on ß-catenin are indirect consequences of the inhibition of cell growth or a direct target of glucocorticoid receptor signaling, Con8 cells were treated with or without 1 µM dexamethasone for 48 h in the presence or absence of 1 mM hydroxyurea. Under these treatment conditions, hydroxyurea abolished the incorporation of [3H]thymidine without affecting cell viability (data not shown), demonstrating that this metabolic agent inhibited DNA synthesis. As shown in Fig. 5AGo, hydroxyurea treatment had no effect on either baseline or dexamethasone-stimulated levels of ß-catenin protein.



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  		Fig. 5. Inhibition of Cell Growth Does Not Regulate ß-Catenin Protein Expression and Localization and Tight Junction Formation

Con8 cells were grown on filter supports, and confluent cells were treated with or without 1 µM dexamethasone (Dex) in the presence or absence of the DNA synthesis inhibitor, 1 mM hydroxyurea (HU) for 48 h. A, Cell lysates were normalized for total protein, electrophoretically fractionated, and transferred onto nitrocellulose membranes. The membranes were probed for ß-catenin and actin protein. B, In a parallel experiment, confluent Con8 cells were grown on the plastic supports and treated with or without 1 mM hydroxyurea for 48 h. Cells were analyzed for ß-catenin and ZO-1 localization by immunofluorescence microscopy. C, To assess the effects of hydroxyurea on tight junction sealing, TER was measured while cells were treated in the presence or absence of hydroxyurea, with or without 1 µM dexamethasone, for 5 d. The figure averages data from three independent experiments.

 
In a parallel experiment, the localization of ß-catenin and ZO-1 proteins was examined by indirect immunofluorescence microscopy. As shown in Fig. 5BGo, addition of hydroxyurea alone had no effect on the nuclear staining of ß-catenin. Although the membrane staining of ZO-1 was somewhat increased in the presence of hydroxyurea, the staining pattern was still punctate and uneven. In the presence of hydroxyurea, dexamethasone treatment increased the membrane staining of both ß-catenin and ZO-1 proteins (Fig. 5BGo, +Dex). In steroid-treated cells, the persistent nuclear staining of ß-catenin, in addition to its membrane localization (Fig. 5BGo, +Dex), is within the levels of experimental variability that we routinely observe between experiments. The results from this experiment demonstrate that the dexamethasone effects on ß-catenin protein level and localization are independent of growth inhibition. To determine whether hydroxyurea exposure has any effect on tight junction sealing, cells were treated with 1 µM dexamethasone and 1 mM hydroxyurea for 5 d, and the monolayer TER was monitored during this time course. As shown in Fig. 5CGo, the results from three independent experiments demonstrate that hydroxyurea treatment had no effect on the kinetics or magnitude of the steroid-induced monolayer TER. Thus, growth inhibition per se has no effect on the glucocorticoid control of ß-catenin expression, junctional organization, and tight junction sealing.

Because ß-catenin protein stability is known to be regulated by GSK3 phosphorylation (34, 35), the effects of dexamethasone treatment on ß-catenin phosphorylation were examined. Con8 mammary tumor cells were treated with or without dexamethasone for 120 h, and the isolated cell extracts were electrophoretically fractionated. The corresponding Western blots were probed with an anti-phospho-ß-catenin antibody that detects phosphorylation at the GSK3-specific sites (serine 33 and 37, and threonine 41) (35), or with an anti-ß-catenin antibody that detects total ß-catenin protein. The production of actin protein was used as a control for gel loading. As shown in Fig. 6AGo, glucocorticoid treatment nearly ablated ß-catenin phosphorylation under conditions in which the level of total ß-catenin protein was increased relative to actin protein levels. Quantification of three independent Western blot experiments showed that steroid treatment significantly reduced specific phospho-ß-catenin by greater than 20-fold (Fig. 6BGo, P < 0.05). This observation suggests that glucocorticoids not only transcriptionally regulate ß-catenin, but may also help stabilize the protein by preventing its phosphorylation. It remains to be determined whether dexamethasone treatment prevents ß-catenin phosphorylation by altering GSK3 signaling or whether the steroid-induced localization of ß-catenin to the cell periphery makes the protein inaccessible to phosphorylation.



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  		Fig. 6. Dexamethasone (Dex) Treatment Ablates Phosphorylation of ß-Catenin at the GSK3 Phosphorylation Sites

A, Con8 cells were treated with and without dexamethasone for 5 d and total cell extracts were electrophoretically fractionated. Western blots were probed with antiphospho (Ser 33/37/ Thr 41) ß-catenin antibodies or anti-ß-catenin antibodies to detect total protein or with antiactin antibodies as a gel loading control. B, Quantification of three independent experiments showed that the 20-fold down-regulation of ß-catenin phosphorylation is statistically significant. Asterisks indicate significance at P < 0.05.

 
TGF{alpha} selectively abrogates the dexamethasone regulation of ß-catenin localization without affecting the steroid-induced increase of ß-catenin protein levels. Our previous work has shown that constitutive expression of TGF{alpha} in transfected cells, or exogenous treatment with TGF{alpha}, prevents glucocorticoid-stimulated tight junction formation in Con8 cells (78, 91). To determine whether TGF{alpha} reverses the effects of dexamethasone on ß-catenin protein levels, CT93 cells, a Con8-derived mammary tumor cell line that constitutively expresses exogenous TGF{alpha}, were treated with or without 1 µM dexamethasone for 5 d. Western blot analysis revealed that dexamethasone was still able to up-regulate ß-catenin protein levels in CT93 cells (Fig. 7AGo) under conditions in which tight junction formation is disrupted (78, 91). Approximately equal loading was confirmed by similar amounts of actin in each sample. ß-Catenin protein levels obtained from three independent experiments in dexamethasone-treated and untreated cells were quantified (Fig. 7BGo). The glucocorticoid-regulated increase in ß-catenin protein levels was statistically significant at the 72-h and the 120-h time points (P < 0.05). The 24-h time point had a P value of only 0.06; however, the observed trend in all experiments was that ß-catenin protein levels were up-regulated in response to steroid treatment. As a complementary approach to examine TGF{alpha} effects on ß-catenin protein production, Con8 cells were treated with or without 1 µM dexamethasone in the presence of 10 ng/ml TGF{alpha} for 5 d, and ß-catenin protein levels were examined by Western blot. As shown in Fig. 7CGo, TGF{alpha} treatment had no effect on dexamethasone up-regulation of ß-catenin protein. Approximately equal loading was confirmed by similar amounts of actin in each sample.



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  		Fig. 7. TGF{alpha} Does Not Alter ß-Catenin Protein Up-Regulation by Dexamethasone (Dex)

A, CT93 cells were seeded on the plastic supports, and confluent cells were treated with or without 1 µM of dexamethasone for 5 d. B, Data from three independent Western blot experiments were quantified with the Scion imaging program and analyzed for statistical significance. Asterisks indicate significance at P < 0.05. C, Confluent Con8 cells were grown on plastic supports and treated with or without 1 µM dexamethasone in the presence of 10 ng/ml TGF{alpha} for 5 d. At each indicated time point, cells were collected and the lysates were normalized for total protein, electrophoretically fractionated, and transferred onto nitrocellulose membranes. The membranes were probed for ß-catenin and actin.

 
For protein localization, a parallel set of cells was treated, fixed, and stained for ß-catenin and ZO-1 by indirect immunofluorescence staining. In CT93 cells (Fig. 8AGo), and Con8 cells treated with 10 ng/ml TGF{alpha} (Fig. 8BGo; +Dex+TGF-{alpha}), dexamethasone failed to increase the membrane staining of ß-catenin and ZO-1 proteins. After 5 d of dexamethasone treatment of CT93 cells and TGF{alpha}-treated Con8 cells, ß-catenin was detected primarily in the cell nucleus, and the ZO-1 staining remained disorganized. Thus, the dexamethasone-regulated localization and expression of ß-catenin can be distinguished by the differential sensitivity to the disruptive effects of TGF{alpha}.



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  		Fig. 8. TGF{alpha} Blocks the Dexamethasone (Dex)-Regulated Localization of ß-Catenin

A, CT93 cells, which express high levels of TGF{alpha}, were seeded on filter supports, and confluent cells were treated with or without 1 µM dexamethasone for 5 d. B, Confluent Con8 cells were grown on filter supports and treated with or without 1 µM dexamethasone in the presence or absence of 10 ng/ml TGF{alpha} for 5 d. At each indicated time point, cells were fixed, and the localization of ß-catenin and ZO-1 proteins was analyzed by immunofluorescence microscopy.

 
Dexamethasone Stimulation of ß-Catenin Protein Expression, But Not Localization, Is a Ras-Dependent Process
Our previous results have shown that Ras-dependent signaling is required for glucocorticoids to enhance the barrier function of the tight junction (79). To determine whether effects of dexamethasone treatment on ß-catenin expression and localization can be distinguished by their requirement for Ras, we used the Con8-derived DN5 cell line, which produces high levels of conditionally inducible dominant negative Ras (RasN17), with significant reduction in Ras GTP levels (79). The RasN17 expression vector was placed under the control of a glucocorticoid-responsive promoter (79). The Con8-derived cell line C7, which was transfected with an empty expression vector, was used as a control for the comparison to the RasN17 transfected cells. Confluent DN5 and C7 cells were treated with or without 1 µM dexamethasone for 5 d, and ß-catenin protein levels were examined by Western blot analysis. As shown in Fig. 9AGo, dexamethasone induced ß-catenin expression level in C7 cells, similar to that seen in Con8 cells (compare to Fig. 1BGo); however, dexamethasone failed to induce ß-catenin protein levels in DN5 cells. Approximately equal loading was confirmed by similar amount of actin in each sample. To assess ß-catenin transcript levels, total RNA was isolated from DN5 cells, treated with or without 1 µM dexamethasone for 24 and 48 h, and analyzed by RT-PCR. As shown in Fig. 9BGo, ß-catenin transcript level remained unchanged after 48 h of steroid treatment, and remained constant in DN5 cells treated for at least 120 h with or without steroid treatment (data not shown). Approximately equal amounts of RNA input was confirmed by the similar amount of GAPDH in each sample.



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  		Fig. 9. Dominant Negative Ras Blocks Dexamethasone (Dex) Up-Regulation of ß-Catenin Protein

RasDN5 and control C7 cells were grown on plastic supports and treated with or without 1 µM dexamethasone for 5 d. At each indicated time point, cell lysates were normalized for total protein, electrophoretically fractionated, and transferred onto nitrocellulose membranes. The membranes were probed for ß-catenin and actin. B, DN5 cells were grown on plastic supports and treated with or without 1 µM dexamethasone for various time points. Total RNA was isolated from each sample, and cDNA was generated by reverse transcription. The transcription level of ß-catenin and GAPDH was amplified by PCR reaction, electrophoretically fractionated on an agarose gel, and visualized by UV light.

 
The localization of ß-catenin and ZO-1 was examined by indirect immunofluorescence staining after 1 d and 5 d with and without dexamethasone treatment. Similar to untransfected Con8 cells, ß-catenin was localized in the nucleus, and ZO-1 staining was disorganized in DN5 cells and C7 cells in the absence of dexamethasone (Fig. 10Go; -DEX). After 5 d of dexamethasone treatment, ß-catenin and ZO-1 proteins were localized to cell-cell contacts and displayed a cobblestone-like staining pattern (Fig. 10Go; +DEX), similar to what we have previously shown for a 4-d treatment with the steroid (79). Compared with untransfected Con8 cells, there was still some detectable ß-catenin localized to the nucleus in both DN5 and C7 cells after 5 d of dexamethasone treatment. Taken together, our results suggest that dexamethasone induction of ß-catenin protein expression, but not the alteration of its localization, is a Ras-dependent process.



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  		Fig. 10. Dominant Negative Ras Does Not Affect ß-Catenin Relocalization by Dexamethasone (Dex)

RasDN5 and control C7 cells were grown on filter supports and treated with or without 1 µM dexamethasone for 5 d. At each indicated time point, cells were fixed and analyzed for ß-catenin and ZO-1 localization by immunofluorescence microscopy.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
ß-Catenin is a multifunctional protein that is not only a major component of adherens junctions but also plays a major role in the Wnt signaling pathway, where it acts as a nuclear transcriptional regulator (41, 82, 83, 84). Thus, the expression and localization of ß-catenin needs to be precisely regulated to ensure cells undergo normal proliferation and differentiation. In our study, we demonstrated that glucocorticoids stimulate ß-catenin expression and nearly ablate GSK3-mediated ß-catenin phosphorylation. Furthermore, steroid treatment facilitates ß-catenin membrane localization as part of the pathway leading to apical junction reorganization and tight junction sealing. As summarized in Fig. 11Go, the dexamethasone stimulation of ß-catenin expression and localization are distinct processes distinguishable by their requirements for Ras and by their differential sensitivity to TGF{alpha} inhibition. The glucocorticoid stimulation of ß-catenin expression can be selectively ablated by overexpression of dominant negative Ras. In contrast, the glucocorticoid induction of ß-catenin localization to the apical junction can be selectively reversed by TGF{alpha}. It has been shown that the formation of the adherens junction in vivo is a prerequisite for the assembly of the tight junction (92, 93). We propose that the multilevel glucocorticoid regulation of ß-catenin is an important step for hormonal stimulation of apical junction formation in mammary epithelial cells and perhaps other differentiated epithelial cells.



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  		Fig. 11. Model for the Effects of Dexamethasone (Dex) on ß-Catenin Protein Expression and Localization

We propose that glucocorticoids regulate ß-catenin at multiple levels. Glucocorticoid up-regulation of ß-catenin protein expression is a Ras-dependent process. TGF{alpha} treatment ablates the glucocorticoid-induced membrane localization of ß-catenin and thus prevents apical junction formation.

 
There is only limited information available on the transcriptional regulation of ß-catenin. Coordinated regulation of the expression of ß-catenin at both mRNA and/or protein levels was reported in cytotrophoblasts (94) hydra bud formation (95), prostate tumor (96), and desmoid tumor (97). However, the molecular mechanisms of regulation were not identified in these studies. In contrast, posttranscriptional regulation of ß-catenin has been well studied and is responsible for many of the dynamic changes that occur during tissue morphogenesis and homeostasis. Using mammary epithelial tumor cells, we observed that GSK3-dependent ß-catenin phosphorylation was nearly ablated after steroid treatment. This observation suggests that glucocorticoids not only transcriptionally regulate ß-catenin, but may also help stabilize the protein by preventing its phosphorylation. It is conceivable that glucocorticoids could directly or indirectly inhibit GSK3 signaling or localize ß-catenin away from its site of phosphorylation. We are currently examining, in more detail, the pathway by which glucocorticoids disrupt ß-catenin phosphorylation.

Oncogenic transformation frequently results in alterations of epithelial properties, including loss of polarized morphology and a decrease in junctional organization. For example, activation of oncogenic Ras proteins in epithelial cells is characterized by mesenchymal/fibroblastic morphology with a perturbation of the adherens junction (98, 99, 100). We observed in Con8 mammary tumor cells that the glucocorticoid-mediated stimulation of ß-catenin protein expression was abrogated by expression of dominant negative RasN17, suggesting that Ras activity is required for dexamethasone induction of ß-catenin. The molecular mechanisms of this process remain elusive. A recent study showed that activated Ras promotes cytoplasmic accumulation of ß-catenin through signaling pathways involving phosphatidylinositol 3-kinase in epidermal keratinocytes (101). Our previous work demonstrated that in Con8 mammary tumor cells, treatment with glucocorticoids recruits Ras and the phosphatidylinositol 3-kinase subunit, p85, to regions of cell-cell contacts, and that the glucocorticoid induction of tight junction sealing is a Ras-dependent process (79). Expression of dominant negative Ras did not abrogate the change in subcellular distribution of ß-catenin, further suggesting that Ras does not play a role in the steroid-mediated reorganization of the apical junction.

Our study demonstrates that dexamethasone induces the localization of ß-catenin to the cell membrane with a concurrent reduction of ß-catenin protein in the nucleus. To date, the molecular mechanisms responsible for the targeting of ß-catenin to the nucleus and to the adherens junction have not been determined. It has been proposed that the different cytoplasmic and nuclear partners of ß-catenin compete with each other for the ß-catenin protein pool and thus determine whether ß-catenin functions in cell adhesion or in transactivation processes (39). Several studies have shown that overexpression of E-cadherin sequesters ß-catenin away from the nucleus and protects ß-catenin from degradation (102, 103), although in the mammary cell system used in our study, dexamethasone treatment had no effect on E-cadherin level. In this regard, there appears to be sufficient E-cadherin and {alpha}-catenin expression in Con8 cells to allow steroid-induced junction formation, thus not requiring additional protein production. Adherens junction formation could also be regulated by protein complex formation or disruption. One study has shown that ß-catenin can bind to cytoplasmic ZO-1 present in MDCK cells cultured in low calcium levels, which leads to tight junction disintegration (104). If a similar process takes places in Con8 cells, a ZO-1/ß-catenin interaction could potentially form a protein complex that prevents both ß-catenin from binding to E-cadherin, and ZO-1 from localizing to the tight junction. Conceivably, dexamethasone treatment may disrupt the formation of a ZO-1/ß-catenin protein complex or perhaps overrides the inhibitory effects of such a protein complex by increasing the level of unassociated ß-catenin protein.

Although nuclear/cytoplasmic shuttling of ß-catenin has been reported before, the mechanisms of ß-catenin nuclear import/export have not been defined. ß-Catenin lacks a classical nuclear localization signal sequence, and one study suggests that ß-catenin may piggyback into the nucleus by binding to T cell factor/Lymphoid enhancing factor or APC (84). Others report that ß-catenin is imported into the nucleus by direct binding to the nuclear pore complex (105). In gonadotropin-releasing neuronal cells, a recent study shows that ligand-bound androgen receptors shuttle ß-catenin to the nucleus (85). The dexamethasone-regulated control of ß-catenin localization demonstrated in our study provides a unique signal-dependent model system to help define the mechanism by which glucocorticoids control ß-catenin nuclear import and export.

Deregulation of ß-catenin signaling has been detected in a number of malignancies, such as colon cancer (106), melanoma (107), hepatocellular carcinoma (108), ovarian cancer (109), endometrial cancer (110), breast cancer (75, 111), and prostate cancer (112). In Con8 mammary tumor cells, glucocorticoids induce normal-like cell-cell interactions that include remodeling of the apical junction and tight junction sealing. In addition, glucocorticoids inhibit Con8 cell growth. Because ß-catenin plays an important role in cell-cell interactions, as well as cell growth and cell survival, investigation of the molecular mechanism of ß-catenin regulation by glucocorticoids will help uncover the pathways that mediate these varied cellular events.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Materials
DMEM/Ham’s F12 [(DMEM/F-12), (50:50)] was supplied by BioWhittaker, Inc. (Walkersville, MD). Permeable tissue culture supports/filter inserts were manufactured by Nunc and distributed by Applied Scientific (San Francisco, CA). Hydroxyurea and RU486 were purchased from Sigma (St. Louis, MO). Human recombinant TGF{alpha} was obtained from Becton Dickinson and Co. (San Jose, CA). Phospho-anti-ß-catenin (Ser33/37/Thr41) antibodies were purchased from Cell Signaling Technology, Beverly, MA). Polyclonal rabbit anti-ZO-1 antibodies and monoclonal mouse anti-ß-catenin antibodies were purchased from Zymed Laboratories, Inc. (South San Francisco, CA). Polyclonal mouse anti-E-cadherin antibodies were purchased from Transduction Laboratories, Inc. (Lexington, KY). Polyclonal rabbit antiactin antibodies were obtained from Sigma. Fluorescein isothiocyanate-conjugated goat antimouse IgG antibodies and Texas red-X conjugated antirabbit IgG antibodies were supplied by Molecular Probes, Inc. (Eugene, OR). Oligonucleotide primers for PCR were purchased from Integrated Diagnostic Technologies (Coralville, IA).

Cell Culture and Measurement of Transepithelial Electrical Resistance
Con8 rat mammary epithelial cells, CT93 cells constitutively expressing TGF{alpha}, C7 cells transfected with a control vector, and DN5 cells expressing the RasN17 dominant negative Ras have been described previously (49, 50, 51, 52). These cell lines were routinely grown to 100% confluency on Nunc permeable supports in DMEM/F-12 supplemented with 10% calf serum and penicillin/streptomycin and maintained at 37 C in a humid atmosphere of air/CO2 (95:5). These cell lines were cultured in serum free medium for 72 h before and during all experiments, and the cell culture medium was routinely changed every 24 h. For the CT93, C7, and DN5 cells, G418 was added into the cell culture medium to a final concentration of 600 µg/ml. In the appropriate experiments, cells were cultured in medium with the synthetic glucocorticoid agonist, dexamethasone (Sigma), or the progesterone/glucocorticoid receptor antagonist RU486 (Sigma) at a final concentration of 5 or 10 µM (prepared as a 1 mM stock in ethanol). Human recombinant TGF{alpha} was added to a final concentration of 10 ng/ml, and hydroxyurea was added to a final concentration of 1 mM. The formation of tight junctions was monitored by measuring TER, using an EVOM Epithelial Voltohmmeter (World Precision Instruments, Sarasota, FL), as described previously (51). Calculations for ohms·cm (2) were determined by subtracting the resistance measurement of a blank filter and multiplying by the area of the monolayer (0.49 cm2 for the 10-mm filters).

Immunofluorescence and Confocal Microscopy
For immunofluorescence analyses, cells were grown on Nunc filters. The cells were washed three times with Dulbecco’s PBS (BioWhittaker, Inc.) and were then fixed with 1.75% formaldehyde in PBS for 15 min at room temperature. After three additional washes with PBS, the plasma membrane was permeabilized with Triton X-100 extraction buffer (0.5% Triton X-100, 100 mM Tris-HCl, pH 7.5, 120 mM NaCl, 20 mM HEPES, and 5 mM EDTA) for 15 min at room temperature. Filters were incubated with a blocking buffer (1% nonfat dry milk in 0.5% Triton X-100, 5 mM EDTA, 0.15 M NaCl, and 20 mM HEPES, pH 7) before incubation with the primary antibodies. Rabbit anti-ZO-1 antibodies and mouse anti-ß-catenin antibodies were used at a 1:400 dilution. Secondary fluorescein isothiocyanate-conjugated antimouse antibodies and Texas red-conjugated antirabbit antibodies were used at a 1:400 dilution. Stained cells were mounted with SlowFade Light Antifade reagent (Molecular Probes, Inc.). Stained and mounted cells were then processed with a Zeiss Axioplan epifluorescence microscope (Carl Zeiss, Thornwood, NY). Confocal fluorescent images of z-y planes were taken with a Zeiss confocal microscope and processed with the corresponding laser scanning microscope software on Microsoft Windows NT at the Berkeley Imaging Center.

Western Blotting
For Western blot analyses, cells were rinsed with PBS (BioWhittaker), and extracted in lysis buffer (50 mM Tris-HPO4, pH 6.8, 2.5 mM EDTA, 15% sucrose, 2% sodium dodecyl sulfate, and 50 mM dithiothreitol) containing protease inhibitors (0.1 M phenylmethylsulfonylfluoride, 1 mg/ml leupeptin, and 1 mg/ml aprotinin). Samples were normalized for protein content with the Bio-Rad Bradford protein assay (Bio-Rad Laboratories, Inc., Hercules, CA). Protein samples were fractionated on a 10% sodium dodecyl sulfate polyacrylamide gel followed by electrophoretic transfer to a nitrocellulose membrane (Micron Separations, Inc., Westborough, MA). Blots were blocked with blotting buffer (5% nonfat dry milk, 0.15 M NaCl, 0.2% Triton X-100) before probing with a 1:1000 dilution of primary antibodies (anti-ZO-1, anti-ß-catenin, antiactin, anti-E-cadherin, and anti-{alpha}-catenin, and antiphospho (Ser 33/37/ Thr 41) ß-catenin). Horseradish peroxidase-conjugated antirabbit and antimouse antibodies (Bio-Rad Laboratories, Inc.) were used as secondary probes, and the blots were developed with an enhanced chemiluminescence kit (Amersham Life Sciences, Inc., Arlington Heights, IL). The level of ß-catenin protein observed in the Western blots was quantified with the Scion imaging program (Scion Corp., Frederick, MD). For comparison of means/t test data analyses the statistical software from Minitab Inc., Release 11 (State College, PA) was used.

RT-PCR
Total RNA from Con8 cells treated with or without 1 µM dexamethasone was isolated with the guanidinium thiocyanate method (95). The DNA in the samples was treated with RQ1 DNase (Promega Corp., Madison, WI). Total RNA (2 µg) in each sample was used to synthesize cDNA using Moloney murine leukemia virus-reverse transcriptase (Promega Corp.) with a random hexamer as a primer in a 20 µl reaction. cDNA reaction product (1 µl) was used with 10 µM ß-catenin primers in the following PCR amplification. The ß-catenin primers are: forward primer, 5'-GATTAACTATCAGGATGACGCG-3'; reverse primer, 5'-TCCATCCCTTCCTGCTTAGTC-3'. As a loading control, 18S rRNA was amplified from the same samples. To adjust the amplification of 18S cDNA, primer pairs for 18S rRNA were mixed with a 1:9 ratio of 18S rRNA competimer pairs (Ambion, Austin, TX), which are primers with a modified 3'-end to block extension. For PCR amplification, 50 µl PCRs (1xTaq polymerase buffer Mg2+ free, 0.2 mM deoxynucleotide triphosphates, 0.25 µl of Taq polymerase (Life Technologies, Inc., Gaithersburg, MD), 1.5 mM MgCl2, 0.2 µM of each primer) were amplified for 24 cycles (95 C, 30 s/55 C, 30 s/68 C, 30 s). The PCRs were precipitated with 1/10 volume of 3 M sodium acetate and 2.5x volume of 100% ethanol, rinsed with 70% ethanol, resuspended in loading dye (32% formamide, 3.3 mM NaOH, 0.05% bromophenol blue, 0.05% xylene cyanol), and separated on a 6% acrylamide gel with 7 M urea, or a 1% agarose gel. To visualize the DNA, the agarose gel was viewed by UV light. In the other case, the acrylamide gel was fixed in 10% acetic acid, stained with 0.1% silver nitrate, 0.06% formaldehyde, and developed with 3% sodium carbonate, 0.06% formaldehyde. For Fig. 3CGo, statistical significance of the increase in ß-catenin transcripts observed in steroid-treated vs. untreated cells was calculated after quantification of silver stain gels from three independent RT-PCR experiments.

Subcellular Fractionation
For fractionation analyses, cells were harvested from tissue culture plates after completion of treatment regimen, lysed for 15 min in hypotonic buffer (10 mM Tris, 10 mM NaCl, 3 mM MgCl2, 1 mM each of EGTA and EDTA), and dounced with 40 strokes in a Dounce homogenizer. Fractions were then spun at 10,000 x g for 10 min. The supernatant was then again spun at 100,000 x g for 60 min to separate the membranes from the cytosol. The nuclear pellet was loaded onto a 2 M sucrose gradient and then spun at 200,000 x g for 60 min at 4C. The subcellular fractionation was verified by probing with the nuclear marker histone deacetylase, and the tight junction protein ZO-1, which is localized to the membranes in steroid-treated cells with functional tight junctions.


   ACKNOWLEDGMENTS
 
This paper is dedicated to the memory of our colleague, Dr. Anita C. Maiyar, whose helpful suggestions throughout most of the study are greatly appreciated. We also thank Bridget A. O’Keeffe for her thoughtful experimental suggestions. We are indebted to Joseph Y. Kim, Joseph Callahan, Daniel H. Lo, Urmi Chatterji, Minnie Wu, Jessie Young, and Sophia Chung for their technical assistance.


   FOOTNOTES
 
This work was supported by NIH Grant DK-42799 (to G.L.F.). P.L.W. was the recipient of a predoctoral fellowship supported by NIH National Research Service Grant CA-09041.

Abbreviations: DAPI, 4,6-Diamidino-2-phenylindole; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GSK3, glycogen synthase kinase 3; TER, transepithelial electrical resistance; ZO-1, zonula occludens-1.

Received for publication January 15, 2003. Accepted for publication October 3, 2003.


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