A Fusion Protein of the Estrogen Receptor (ER) and Nuclear Receptor Corepressor (NCoR) Strongly Inhibits Estrogen-Dependent Responses in Breast Cancer Cells

Pei-Yu Chien, Masafumi Ito, Youngkyu Park, Tetsuya Tagami, Barry D. Gehm and J. Larry Jameson

Division of Endocrinology, Metabolism, and Molecular Medicine Northwestern University Medical School Chicago, Illinois 60611


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Nuclear receptor corepressor (NCoR) mediates repression (silencing) of basal gene transcription by nuclear receptors for thyroid hormone and retinoic acid. The goal of this study was to create novel estrogen receptor (ER) mutants by fusing transferable repressor domains from the N-terminal region of NCoR to a functional ER fragment. Three chimeric NCoR-ER proteins were created and shown to lack transcriptional activity. These fusion proteins silenced basal transcription of the ERE2-tk-Luc reporter gene and inhibited the activity of cotransfected wild-type ER (wtER), indicating that they possess dominant negative activity. One of the fusion proteins (CDE-RD1), containing the ER DNA-binding and ligand-binding domains linked to the NCoR repressor domain (RD1), was selected for detailed examination. Its hormone affinity, intracellular localization, and level of expression in transfected cells were similar to wtER, and it bound to the estrogen response element (ERE) DNA in gel shift assays. Glutathione-S-transferase pull-down assays showed that CDE-RD1 retains the ability to bind to steroid receptor coactivator-1. Introduction of a DNA-binding domain mutation into the CDE-RD1 fusion protein eliminated silencing and dominant negative activity. Thus, the RD1 repressor domain prevents transcriptional activation despite the apparent ability of CDE-RD1 to bind DNA, ligand, and coactivators. Transcriptional silencing was incompletely reversed by trichostatin A, suggesting a histone deacetylase-independent mechanism for repression. CDE-RD1 inhibited ER-mediated transcription in T47D and MDA-MB-231 breast cancer cells and repressed the growth of T47D cells when delivered to the cells by a retroviral vector. These ER-NCoR fusion proteins provide a novel means for inhibiting ER-mediated cellular responses, and analogous strategies could be used to create dominant negative mutants of other transcription factors.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The estrogen receptor (ER) belongs to the superfamily of nuclear receptors, which regulate the transcription of specific target genes. The ER binds to the estrogen response element (ERE), a palindromic DNA sequence in the promoters of estrogen-regulated genes (1, 2), and activates transcription in response to its ligand, estradiol (E2). Several functional domains have been identified in the ER (3), including transactivation domain 1 (TAF-1, regions A/B), a DNA-binding domain (region C), a hinge region (region D), and transactivation domain 2 (TAF-2)/hormone-binding domain (region E). In most cases, both TAF-1 and TAF-2 are required for full hormone-stimulated activity. It has been shown recently that there are two different isoforms of ER, ER{alpha} (4, 5) and ERß (6, 7), that are encoded by different genes. These two isoforms are highly similar in the DNA-binding domain and, to a lesser degree, in the ligand-binding domain (6, 7).

Hormone-induced gene transcription can be enhanced by coactivators, including steroid receptor coactivator-1 (SRC-1) (8, 9), SRC-2 [also called TIF2 (transcriptional intermediary factor) and GRIP-1 (glucocorticoid receptor interacting protein 1 ) (10, 11)], SRC-3 [also called p/CIP (p300/CBP cointegrater- associated protein), RAC3 (receptor-associated coactivator 3), ACTR (activator of thyroid receptor), and AIB1) (amplified in breast cancer 1) (12, 13, 14, 15), and CREB binding protein (CBP)/p300 (16, 17), among others. Recently determined crystal structures of nuclear receptors reveal that helices 3, 5, and 12 of the ligand binding domain form a hydrophobic groove that serves as the interaction surface for an hydrophobic {alpha}-helical segment (LXXLL) in coactivators (18, 19). A translocation of helix 12 is induced by ligand binding and plays a critical role in the recruitment of coactivators (18). Therefore, recruitment of coactivators to TAF-2 of nuclear receptors is responsible for ligand-dependent gene activation. Coactivators also interact with the N-terminal (TAF-1) region of nuclear receptors (20, 21), and this region contributes to ligand-independent gene activation (22). SRC-1 increases ER transcriptional activity by enhancing the interaction between the TAF-1 and TAF-2 domains of ER (23). In addition, coexpression of CBP/p300 and SRC-1 synergistically stimulates ER transcriptional activity (24). The conformational changes induced by ligand binding to ER recruit a coactivator complex containing SRC-1, CBP/p300, and p300/CBP-associated factor (P/CAF) (25). These coactivators have recently been found to have intrinsic histone acetyltransferase (HAT) activity (12, 26, 27, 28). HAT acetylates histones, a process that is proposed to alter the chromatin structure and increase the accessibility of DNA to transcription factors and the basal transcription machinery (29, 30), thereby enhancing the rate of gene transcription (31). HAT also acetylates components of the basal transcription complex (32), in addition to histones.

On the other hand, in the absence of ligand, several nuclear receptors suppress basal gene transcription by recruiting corepressors (CoRs). Nuclear receptor corepressor (NCoR) (33) and silencing mediator for retinoic acid and thyroid hormone receptors (SMRT) (34, 35) have been intensively studied in the context of the unliganded thyroid hormone receptor (TR) and retinoic acid receptor (RAR). The amino acid sequences of NCoR and SMRT show approximately 40% identity (35). NCoR contains three transferable repressor domains in its N-terminal region (36), of which the first (RD1) appears to be the most potent (33). SMRT is a shorter protein and has two repressor domains in its N-terminal region, which corresponds to the third repressor domain of NCoR (37). Unliganded TR and RAR recruit a multicomponent CoR complex which, in addition to NCoR or SMRT, also contains mSin3, a mammalian homolog of a yeast transcription CoR (38, 39), and histone deacetylase (HDAC) (36, 37). HDAC deacetylates histone H3 (40, 41), stabilizing the structure of chromatin, and inducing repression of basal gene transcription. Based on these recent findings, it is believed that chromatin remodeling is a key event in the regulation of gene transcription by nuclear receptors and other transcription factors (reviewed in Refs. 42, 43, 44, 45, 46).

In contrast to TR and RAR, suppression of ER-mediated basal transcription by CoRs has not been observed. However, there is evidence that ER interacts with CoRs in vitro. In glutathione-S-transferase (GST) fusion pull-down assays, SMRT binds to ER in a hormone-independent manner (47). NCoR coimmunoprecipitates with ER weakly without ligand and more strongly in the presence of tamoxifen (48), an ER-mixed antagonist/agonist (49, 50).

Dominant negative mutants of ER, which block the function of the wild-type ER (wtER), have been created by chemical or site-directed mutagenesis (51, 52, 53, 54). These mutants retain DNA binding and dimerization properties, but they are deficient in transcriptional activation, allowing them to function as antagonists at ERE binding sites. As the RD1 of NCoR retains its repressor activity when fused to DNA binding proteins such as Gal4 (33), we hypothesized that dominant negative variants of ER could be produced by fusing the NCoR repressor domains to the ER.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
A schematic diagram of the proteins used in this study is shown in Fig. 1Go. The wild-type ER contains six functional domains (A to F). The CDE fragment used in this study is a truncated form that contains the DNA binding domain, hinge region, and TAF-2/ligand binding domain. NCoR contains repressor domains 1, 2, and 3 (RD1, RD2, and RD3) in its N-terminal region (36) and two C-terminal interaction domains (ID1 and ID2), which interact with the ligand binding domain of nuclear receptors like TR and RAR (33, 55). RD1 was fused to the C terminus of CDE to create CDE-RD1 and to the N terminus to create RD1-CDE. The whole repressor region of NCoR, containing all three of the repressor domains, was fused to the N terminus of CDE to create RD1–3-CDE.



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Figure 1. Schematic Diagram of the wtER, NCoR, and Fusion Proteins

The functional domains of wtER (A–F) and NCoR (RD1, RD2, RD3, ID1, and ID2) are displayed. CDE, a truncated form of ER, contains the DNA-binding domain (C), hinge region (D), and TAF-2/ligand binding domain (E). Repressor domain 1 (RD1) of NCoR, the most potent repressor domain, was fused to the C terminus of CDE to create CDE-RD1 or to the N terminus to create RD1-CDE. The whole repression region (RD1–3) was fused to the N terminus of CDE to create RD1–3-CDE. The amino acid residues corresponding to functional domains are indicated.

 
Effect of Repressor Domain(s) on the Transcriptional Activity of Fusion Proteins
The transcriptional effects of various ER constructs were examined using the estrogen-responsive reporter, ERE2-tk109-luc, in ER-negative TSA-201 cells (Fig. 2Go). In the absence of transfected ER, E2 did not stimulate the reporter activity in these cells. Cotransfection with an expression vector encoding wtER conferred 10-fold estrogen responsiveness. Control transfections with empty expression vectors, or with plasmids containing unrelated proteins, did not alter estrogen responsiveness, either in the absence or presence of the wtER (data not shown). The truncated ER variant, CDE, also conferred estrogen responsiveness, but the E2-stimulated reporter activity was about half that obtained with wtER. The activity of CDE presumably represents transcription stimulated by the TAF-2 domain in the absence of TAF-1. In contrast, the fusion protein CDE-RD1 showed no estrogen-induced stimulation of reporter gene expression. RD1-CDE and RD1–3-CDE were also deficient in E2 responsiveness (data not shown). These data suggest that the fusion proteins are not expressed, are defective in E2 binding, or that the repressor domain(s) inhibit the activity of TAF-2, resulting in the loss of E2-induced transcriptional activity. CDE-RD1 was chosen for more detailed characterization (see Discussion).



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Figure 2. Fusion Protein CDE-RD1 Is Transcriptionally Inactive

TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc reporter plasmid and 50 ng/well of control vector (pCMX-CAT), pCMX-ER (wtER), pCMX-CDE, or pCMX-CDE-RD1. After transfection, cells were treated and assayed as described in Materials and Methods. Results of all luciferase assays are shown as the mean ± SD of quadruplicate transfections.

 
Expression of CDE-RD1
The extreme N-terminal domain of NCoR has been shown to target NCoR for proteasomal degradation (56). Therefore, the N-terminal repressor domain might target the fusion protein for degradation, resulting in poor expression. To examine this possibility, the presence of CDE-RD1 in transfected cells was confirmed using immunocytochemistry and Western blotting. Immunostaining using the anti-ER antibody H222 showed that wtER was expressed in the nuclei of TSA-201 cells transfected with pCMX-ER (Fig. 3AGo). Both CDE and CDE-RD1 were also expressed in the nuclei of the transfected cells (Fig. 3Go, B and C). In contrast, no staining was detected in cells transfected with empty vector (Fig. 3DGo) or when control rat IgG was used in place of H222 (data not shown).



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Figure 3. CDE-RD1 Is Expressed in Transfected Cells

A–D, TSA-201 cells were transfected and subjected to immunostaining as described in Materials and Methods. Photomicrographs are shown at 400x magnification. Staining is visible mainly in nuclei, as indicated by arrowheads. E, Western blot. Nuclear extracts were prepared, blotted, and probed as described in Materials and Methods. Lanes 1–4 represent cells transfected with no ER, wtER, CDE, and CDE-RD1, respectively. The scale at left indicates the position and molecular mass (kDa) of marker proteins.

 
Nuclear extracts of transfected TSA-201 cells were also analyzed by Western blot (Fig. 3EGo). Antibody H222 revealed similar levels of wtER and CDE-RD1 (lanes 2 and 4) and a greater level of expression of CDE (lane 3). The mobilities of the wtER, CDE, and CDE-RD1 bands are consistent with their predicted molecular masses of 66, 45, and 82 kDa, respectively. Two faint bands of smaller molecular mass were detected in lanes 2 and 4. These bands may result from proteolytic degradation of wtER (lane 2) and CDE-RD1 (lane 4). No ER expression could be detected when the cells were transfected with empty vector (lane 1). The immunostaining and Western blotting demonstrate that the expression and localization of CDE-RD1 in transfected TSA-201 cells are similar to that of wtER. Thus, lack of CDE-RD1 activity is not simply due to degradation.

Hormone Binding to CDE-RD1
Although CDE-RD1 contains the ER ligand-binding domain, it is possible that the attachment of RD1 could interfere with estrogen binding, accounting for the loss of transcriptional activity. We therefore examined the affinity of the fusion protein for estrogen. CDE-RD1 was synthesized by in vitro transcription/translation, and Scatchard analysis of 125I-estradiol binding yielded a Kd of 0.38 nM (data not shown), which is comparable to that of the wtER (0.5 nM) (57). Thus, the absence of transcriptional activity by CDE-RD1 is not due to loss of hormone binding.

DNA binding of CDE-RD1
The inactivity of CDE-RD1 could also be caused by loss of DNA binding. The ability of CDE-RD1 to bind to the ERE in vitro was examined using electrophoretic mobility shift assays. Nuclear extracts from TSA-201 cells transfected with expression vectors for wtER, CDE, CDE-RD1, or no ER (empty vector) were incubated with a radiolabeled ERE probe (Fig. 4Go). The binding of wtER to the ERE probe is shown in lane 2. The CDE fragment also binds to the ERE and exhibits more rapid migration, consistent with the lower molecular mass of CDE (lane 3). Conversely, the larger CDE-RD1 protein produced a more slowly migrating band (lane 4), which was also less intense. Preincubation with anti-ER antibody (AER308) supershifted the wtER, CDE, and CDE-RD1 bands, confirming the identity of the proteins (data not shown). The bands were also eliminated by a 100-fold excess of nonradioactive probe, indicating specific binding to the ERE (data not shown). These and other experiments confirm that CDE-RD1 binds to DNA, although the amount of binding was consistently reduced, suggesting weaker affinity, or reduced stability, of the protein-DNA complex.



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Figure 4. CDE-RD1 Binds to the ERE

Nuclear extracts from TSA-201 cells transfected with empty vector (no ER), wtER, CDE, or CDE-RD1 were incubated with 32P-labeled ERE and analyzed by PAGE as described in Materials and Methods.

 
Coactivator Binding to CDE-RD1
Because the ligand binding domain of ER is fully intact in CDE-RD1, the fusion protein might still be able to bind SRC-1. We examined this possibility using GST pull-down assays. GST was fused to the central region [amino acid (a.a.) 661 to a.a. 855] of SRC-1, which contains three LXXLL (where L is leucine and X is any a.a.) motifs (also called NR boxes) that are important for binding to nuclear receptors, including ER (11, 21, 58, 59, 60). 35S-labeled wtER or CDE-RD1 was incubated with GST, or with the GST-SRC-1 fusion protein, in the presence or absence of E2 (Fig. 5Go). Unmodified GST bound very little wtER or CDE-RD1. In contrast, GST-SRC-1 bound wtER in an E2-dependent manner, reflecting hormone-dependent association of the receptor and coactivator. Similar results were obtained with CDE-RD1, indicating that the presence of the repressor domain does not preclude binding to SRC-1. Thus, CDE-RD1 is transcriptionally inactive despite its ability to bind DNA, ligand, and coactivators.



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Figure 5. CDE-RD1 Interacts with GST-SRC-1 in Vitro

35S-labeled wtER (upper panel) or CDE-RD1 (lower panel) were applied to glutathione-agarose beads pretreated with bacterially expressed GST or GST-SRC-1 in the presence or absence of 10 nM E2. Bound proteins were eluted, resolved by 10% SDS-PAGE, and visualized by autoradiography.

 
Inhibition of wtER Transcriptional Activity by ER-NCoR Fusions
All three fusion proteins (CDE-RD1, RD1-CDE, and RD1–3-CDE) were examined for their abilities to inhibit the activity of wtER. ER-deficient TSA-201 cells were transfected with an estrogen-responsive reporter plasmid, a constant amount of wtER expression vector, and varying amounts of the fusion protein vectors, such that the ratio of chimera/wt ER ranged from 0.1 to 10. After transfection, the cells were treated with or without E2. As shown in Fig. 6AGo, each of the fusion proteins inhibited wtER activity in a dose-dependent manner. In cells transfected with a 10-fold excess of CDE-RD1 or RD1-CDE vectors, E2-induced transcription was almost completely (~98%) inhibited. RD1–3-CDE produced substantial inhibition, but it was not as potent as the other two fusion proteins. When the reporter gene was replaced with tk109-luc (the same reporter without an ERE), E2 did not increase transcription, and the activity of the reporter was not affected by the addition of the fusion proteins (data not shown). These data indicate that fusion proteins of RD1 and CDE have a strong dominant negative effect with respect to the wtER and that the dominant negative effect is dependent on the presence of the ERE.



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Figure 6. Repressor Fusion Proteins Block wtER Activity, and This Dominant Negative Activity Requires DNA Binding

TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc, 5 ng/well pCMX-ER, and 0.5 to 50 ng/well of fusion protein expression vectors in panel A or 5 or 50 ng/well of DNA binding mutant in panel B, resulting in the indicated range of fusion/wt vector ratios. Empty vector was added to provide equal amounts of total DNA in each well. C, The dominant negative effect of CDE-RD1 is specific for ER. TSA-201 cells were transfected with 100 ng/well GRE-tk109-luc, 5 ng/well RSV-GR, and 50 ng/well empty vector (pCMX), wtER, or CDE-RD1. Six hours after transfection, cells were treated with 100 nM dexmethasone or control vehicle and assayed for luciferase activity 48 h later.

 
To further investigate whether the dominant negative activity of CDE-RD1 requires DNA binding, a DNA binding-deficient version of CDE-RD1 was created by introducing three point mutations (E203G, G204S, and A207V) into the C region of CDE-RD1. These three amino acids in nuclear receptors have been shown to play a key role in DNA binding by the ER (61, 62). The introduction of the DNA-binding domain mutations into CDE-RD1 eliminated ERE binding in electrophoretic mobility shift assays, but did not affect the expression level of the fusion protein in Western blot (data not shown). In transient transfection assays, when wtER was cotransfected with CDE-RD1 at a ratio of 1:1, the wtER activity was strongly inhibited, whereas no repression was observed when wtER was cotransfected with DNA-binding mutant (Fig. 6BGo). These results confirm that full dominant negative activity by the fusion protein is dependent on DNA binding. Notably, the DNA binding-deficient mutant produced some inhibition when transfected at 10-fold excess over wtER. This inhibition may be mediated by protein-protein interactions. Mutating the DNA-binding domain also abolished the basal silencing effect of the fusion protein (data not shown).

The effect of CDE-RD1 was also tested with respect to glucocorticoid receptor (GR)-mediated transcription to examine whether its inhibitory activity was specific for the ER, or also occurred with other steroid receptors. In TSA-201 cells transfected with RSV-GR (a GR expression vector) and a glucocorticoid-responsive reporter (GRE-tk109-luc), treatment with dexamethasone induced a 3.8-fold increase in luciferase activity (Fig. 6CGo). Cotransfection of wtER expression vector with GR at a 10:1 ratio reduced (37%), but did not eliminate, the response to dexamethasone. This inhibition may be due to competition for transcriptional cofactors (squelching). In contrast, cotransfection with CDE-RD1 expression vector did not inhibit dexamethasone-induced activity, and dexamethasone induced a 4.2-fold increase of reporter activity. Thus, CDE-RD1 does not suppress GR-mediated transcription, suggesting that its dominant negative effect is specific for ER.

Comparison of CDE-RD1 with Other Dominant Negative ER Mutants
Several dominant negative ER mutants have previously been created by chemical and site-directed mutagenesis (52, 53, 54). We compared the dominant negative activity of CDE-RD1 to two of these mutants: L540Q, a point mutation in the TAF-2 domain, and ER1–536, a C-terminal truncation of the TAF-2 domain. TSA-201 cells were cotransfected with the ERE2-tk109-luc reporter plasmid, wtER, and expression vectors for L540Q, ER1–536, or CDE-RD1 (at ratios to the wtER ranging from 1 to 10) (Fig. 7Go). At a ratio of 1:1, CDE-RD1 exhibited about twice the inhibitory activity of either L540Q or ER1–536. At a ratio of 10:1, CDE-RD1 completely blocked wtER activity, whereas the inhibition by L540Q and ER1–536 was partial.



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Figure 7. Comparison of CDE-RD1 with Other Dominant Negative ER Mutants

TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc, 5 ng/well of pCMX-ER, and 5–50 ng/well of expression vectors for L540Q, ER1–536, or CDE-RD1 producing mutant/wt ratios ranging from 1 to 10, as indicated. Empty vector was added to make the total DNA in each well equal. Cells were treated and assayed as described in Materials and Methods.

 
Inhibition of ER-Dependent Responses in Breast Cancer Cells
Since dominant negative ER mutants might be useful for inhibiting estrogen action in breast cancer cells, we examined the dominant negative effect of CDE-RD1 in two breast cancer cell lines. ER-negative MDA-MB-231 cells were cotransfected with wtER and CDE-RD1. At a ratio of 1:1, E2 induction of wtER activity (assessed by the reporter ERE2-tk109-luc) was inhibited by 50%, and a 10:1 ratio produced 85% inhibition (Fig. 8AGo). In ER-positive T47D breast cancer cells, CDE-RD1 inhibited endogenous ER activity in a dose-dependent manner (Fig. 8BGo). Using 100 ng of CDE-RD1 expression vector, estrogen-stimulated reporter activity was decreased by 70%. Inhibition was almost complete (>90%) using 500 ng of the CDE-RD1 expression vector. These data show that CDE-RD1 exhibits a dominant negative effect with respect to endogenous receptor in breast cancer cells.



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Figure 8. CDE-RD1 Inhibits wtER Transcriptional Activity in Breast Cancer Cells

A, MDA-MB-231 (ER-negative) breast cancer cells were transfected with 1 µg/well ERE2-tk109-luc, 5 ng/well of wtER, and 5 ng/well or 50 ng/well of CDE-RD1 expression vectors resulting in 1:1 or 10:1 CDE-RD1/wt ratios. After transfection, cells were treated and assayed as described in Materials and Methods. B, T47D (ER-positive) breast cancer cells were transfected with the ERE2-tk109-luc (2 µg/well) and the CDE-RD1 expression vector (0–500 ng/well). Forty-eight hours after transfection, cells were treated with 1 nM E2 or control vehicle and assayed for luciferase activity.

 
To study the effect of CDE-RD1 on estrogen-stimulated growth of breast cancer cells, a retroviral delivery system was used to achieve more quantitative introduction of the mutant into the cells. T47D cells were transduced with medium collected from packaging cells transfected with empty (control) or CDE-RD1-containing retroviral vectors and assayed for cell growth. CDE-RD1 mRNA was detected by RT-PCR in cells transduced with CDE-RD1 retrovirus but not with empty retrovirus (data not shown), confirming delivery of the CDE-RD1 gene. As shown in Fig. 9Go, at assay day 7, the cells transduced with control retrovirus grew well in response to E2 (>400% increase over day 1). In contrast, the cells transduced with CDE-RD1 retrovirus proliferated significantly slower in the presence of E2 (90% increase over day 1) and did not grow at all in the absence of E2.



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Figure 9. CDE-RD1 Suppresses the Growth of Breast Cancer Cells

T47D cells were transduced with control or CDE-RD1-containing retrovirus as described in Materials and Methods. The cells were treated with 1 nM E2 or control vehicle, and the cell density was determined at the indicated intervals. Each point represents the mean ± SD of five replicate wells.

 
Silencing Effect of CDE-RD1
Unliganded TR and RAR repress or silence basal gene transcription via interaction with the NCoR complex (33, 36, 55, 63). Since transcriptional silencing is mediated by the repressor domains of NCoR, we tested whether CDE-RD1 might also repress basal transcription of ER-responsive genes. The activity of the ERE2-tk109-luc reporter gene in TSA-201 cells transfected with control vector (100 ng) and treated with control vehicle (ethanol) established the level of basal expression (indicated by dashed line in Fig. 10AGo). Transfection with ER (100 ng) produced higher reporter activity, even in the absence of E2. This may represent transcriptional activity by the unliganded receptor (64), or it may be due to traces of estrogen, even though the cells were cultured in estrogen-depleted medium for 4 days. In contrast, transfection with CDE-RD1 decreased reporter activity below the basal level. This suppression was dose-dependent, with 100 ng of expression vector producing a 70% decrease in activity (Fig. 10AGo). When CDE-RD1 was cotransfected with ER, the unliganded ER activity was also inhibited by CDE-RD1 (Fig. 10BGo). This basal inhibitory effect was not detected when tk109-luc was used instead of ERE2-tk109-luc (data not shown), indicating that it requires the presence of the ERE. Thus, incorporation of RD1 into ER produces a protein that silences basal reporter gene transcription and unliganded wtER activity.



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Figure 10. CDE-RD1 Silences Basal and Unliganded wtER Gene Transcription

A, TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc and 100 ng of control or wtER expression vectors, or 1, 10, or 100 ng of pCMX-CDE-RD1. Basal activity is indicated by a dashed line. B, TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc and 5 ng of control or wtER expression vectors, or 5 ng of wtER expression vector plus 50 ng of pCMX-CDE-RD1. Six hours after transfection, fresh estrogen-depleted medium was added and cells were assayed as described in Materials and Methods.

 
Mechanism of Silencing by CDE-RD1
As previously described, NCoR recruits HDAC (36), which inhibits the transcription of many promoters. We used the HDAC inhibitor, trichostatin A (65, 66, 67, 68, 69), to examine the role of HDAC in the action of CDE-RD1. TSA-201 cells were transfected with the ERE2-tk109-luc reporter plasmid and control vector, wtER, or CDE-RD1, as shown in Fig. 11Go. In the absence of ER (control), the reporter activity was unaffected by E2, and the fold induction by trichostatin A was similar in the unliganded and E2-treated groups (23- and 27-fold, respectively). This induction suggests that the reporter construct is sensitive to trichostatin A, independent of the ER. In cells transfected with wtER, both ligand-independent and E2-stimulated activity were strongly increased by trichostatin A, but the fold increase was less for the E2-treated group (14-fold vs. 27-fold). In cells transfected with CDE-RD1, trichostatin A increased basal transcription, but the fold increase was smaller than that in control cells, and E2 produced no further induction (15-fold and 11-fold for the unliganded and E2-treated groups, respectively). The inability of trichostatin A to completely reverse silencing by fusion protein RD1 suggests that the inhibition by RD1 may involve an HDAC1-independent mechanism.



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Figure 11. Effect of the HDAC1 Inhibitor, Trichostatin A, on Transcriptional Silencing

TSA-201 cells were transfected with 500 ng/well ERE2-tk109-luc and 5 ng/well of control, or wtER, or 55 ng CDE-RD1 expression vectors. Cells were treated with or without 1 nM E2 and with or without 100 nM trichostatin A. Results are shown as the mean ± SD of quadruplicate transfections. Inset, Same data plotted on expanded scale to facilitate comparison of different ER regimens in the absence of trichostatin A.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Several dominant negative ER mutants have been created previously by chemical or site-directed mutagenesis (51, 52, 53, 54). These mutants inhibit ER stimulation of target genes, presumably by competing for binding to the ERE, or by forming inactive dimers with the wtER. Recently, studies of dominant negative mutants of the TR revealed an unanticipated role for CoRs in their inhibitory activity (70, 71). For example, mutations in the TR that specifically disrupt binding to NCoR or SMRT almost completely eliminate dominant negative activity of naturally occurring mutants that cause resistance to thyroid hormone (70, 72). These features suggest that the recruitment of CoRs is an important component of inhibition by these mutants. Because the wtER interacts relatively weakly with CoRs (47, 48), we reasoned that the dominant negative activity of ER mutants might be enhanced if they were fused to CoRs.

Three different fusion proteins between ER and NCoR were created in an effort to identify a configuration that would retain ER properties such as dimerization and DNA binding, but allow the repressor domains to contribute to transcriptional inhibition. The smaller proteins, CDE-RD1 and RD1-CDE, contained one of the repressor domains (RD1) of NCoR fused to either the carboxy terminus or the amino terminus of a truncated form of the ER (CDE). In another mutant, all three repressor domains (RD1–3) were fused to the amino terminus. The CDE region of the ER was used, rather than the full-length ER, for several reasons. Elimination of the TAF-1 domain (A/B region) was anticipated to reduce the constitutive transcriptional activity conferred by this region (3, 73). As shown in Fig. 2Go, CDE retained about half of the transcriptional activity of the wtER. These experiments also suggest that this truncated form of the receptor retains essential functions of the ER, such as estrogen binding and the ability to bind and activate target genes. We also eliminated the F domain from these constructs. Although the function of this domain is not fully understood, it may play a role in functional differences mediated by agonists and antagonists (74, 75). Deletion of the A/B and F domains permitted creation of fusion proteins with molecular masses similar to that of wtER. The predicted molecular mass for the fusion protein with RD1 is 82 kDa, which is close to that of wtER (66 kDa). In contrast, the predicted size of the RD1–3-CDE fusion protein is much greater (233 kDa). In electrophoretic mobility shift assays, CDE-RD1 (Fig. 4Go) and RD1-CDE (data not shown) retained dimerization and ERE binding, whereas little or no DNA binding was seen with the larger RD1–3-CDE fusion protein (data not shown). For this reason, we concentrated the studies on CDE-RD1, which also retained E2 binding, nuclear targeting, and a near-normal level of protein expression (Fig. 3Go). It is possible that the larger fusion protein (RD1–3-CDE) blocks ERE binding or is unstable under conditions of the gel shift assay.

The addition of the RD1 domain suppressed the activity of the truncated ER, in both the absence and presence of E2. Current models hold that the binding of ligand induces conformational changes in nuclear receptors that release the repressor complex (for receptors like TR or RAR) and allow the recruitment of a coactivator (CoA) complex that stimulates gene transcription. In the ER, helices 3, 5, and 12 (18, 76, 77) and several residues in the ligand-binding domain, such as Leu417 and Glu420 (78), are important for the interaction with the coactivator, SRC-1. These regions of the ER are intact in CDE, which retains ligand-dependent transcriptional activation (Fig. 2Go). Therefore, it is intriguing that the repressor domain of NCoR is sufficient to suppress the transcriptional activity of the TAF-2 domain. One possible explanation is that the repressor domain, which cannot be released from the fusion protein, precludes the recruitment of SRC-1. However, the GST pull-down assay demonstrated that CDE-RD1 retains interactions with SRC-1 (Fig. 5Go). Since GST pull-down assay is a simplified system and can only reflect in vitro behavior, we cannot rule out the possibility that the repressor domain might still preclude the recruitment of SRC-1 or other components of the CoA complex in vivo. Based on reporter gene assays, even if SRC-1 is recruited to CDE-RD1 through a ligand-induced conformation change, the TAF-2 domain remains inactive, indicating that the repressor domain might suppress the TAF-2 activity by inhibiting CoA function. Thus, the repressor complex may inhibit transcription so strongly that it must be removed before the HAT activity of the CoA complex can become effective.

Consistent with the irreversible inactivation of the ER by the repressor domain, fusion proteins such as CDE-RD1 are also effective as dominant negative inhibitors of wtER. CDE-RD1 inhibits wtER activity under both liganded (Fig. 6AGo) and unliganded conditions (Fig. 10BGo). CDE-RD1 did not inhibit GR-mediated transcription (Fig. 6CGo), indicating that its dominant negative effect is specific for the ER. When compared with other dominant negative ER mutants, such as L540Q (a point mutation) and ER1–536 (a carboxy-terminal truncation), CDE-RD1 appeared to exert more potent inhibitory activity (Fig. 7Go). It is notable, however, that the ER dominant negative mutants are reasonably effective in the absence of the RD1 fusion protein. It is possible that these mutants interact with cellular CoRs more effectively than the unliganded wtER, but this remains to be investigated. These previously reported dominant negative mutants of ER result from mutation or deletion in helix 12 of the TAF-2 domain, causing loss of the ability to recruit coactivators. Our data indicate that CDE-RD1 can interact with SRC-1 and so represents a novel class of dominant negative ER mutants.

Given the strong dominant negative activity of CDE-RD1, we decided to further explore its effectiveness using breast cancer cell lines. As shown in Fig. 8Go, CDE-RD1 inhibited both exogenous and endogenous ER activity in this model. Furthermore, using retroviral delivery of CDE-RD1, we were able to demonstrate significant growth suppression of these breast cancer cells (Fig. 9Go). Combined with another recent study using adenovirus to deliver dominant negative ER mutants into breast cancer cells (79), these data suggest a potential utility for targeting ER in the treatment of breast cancer.

In addition to its ability to block E2-mediated transcription, CDE-RD1 also suppressed basal gene transcription in a manner reminiscent of that seen with unliganded TR and RAR (80, 81, 82) (Fig. 10AGo). Transcriptional silencing by unliganded TR and RAR involves the recruitment of CoRs such as NCoR and SMRT, which in turn recruit a complex containing Sin3 and HDAC (36). HDAC deacetylates histone, a process that is proposed to stabilize chromatin structure and thereby suppress gene transcription (reviewed in Refs. 42, 43, 44, 45, 46). Regions within NCoR that have been shown to mediate the interaction with Sin3 include an amino-terminal SIN interaction domain (SID 1, a.a. 254–312) and a second domain (SID 2, a.a. 1829–1940) (36, 63). The RD1 (a.a. 1–312) fragment studied here includes SID1. Therefore, fusing RD1 with ER might be expected to recruit SIN3 and suppress gene transcription by a mechanism that involves the recruitment of HDAC.

Based on this idea, we used trichostatin A, an inhibitor of HDAC1, to assess its effect on transcriptional suppression by CDE-RD1 (Fig. 11Go). Trichostatin A increased the activity of the reporter gene, even under control conditions, suggesting that the promoter is repressed under basal conditions by histone deacetylation. However, the effect of trichostatin A on ERE2-tk109-luc activity was much greater in the presence of E2-stimulated ER, indicating that inhibition of histone deacetylation enhances transcriptional activation by the ER (83, 84). Unexpectedly, trichostatin A did not reverse the inhibitory effect of the repressor domain in CDE-RD1. Similar findings have been reported for suppression by the retinoblastoma protein (Rb). Rb is a tumor suppressor protein that inhibits gene transcription, in part through interactions with E2F, a family of cell cycle transcription factors (85). Rb has been shown to suppress gene transcription through both HDAC-dependent (69, 86, 87) and HDAC-independent (87, 88) mechanisms. The amino-terminal region of NCoR (a.a. 1–1017) has been shown to physically interact with basal transcription factors in vivo and in vitro, and NCoR can inhibit the recruitment of the TAFII-32 subunit of TFIID by TFIIB (89). RD1 (a.a. 1–312) used in our study is included within this amino-terminal region. Therefore, it is possible that CDE-RD1 might silence gene transcription through interactions with the basal transcriptional machinery. CDE-RD1 might also recruit an unidentified HDAC that is resistant to inhibition by trichostatin A.

In summary, we have demonstrated that fusing a repressor domain from NCoR to a truncated form of ER can irreversibly silence ER function and inhibit wtER activity when cotransfected into cells. This strategy offers a new approach for the study of estrogen action and, with modification, might be useful for gene therapy for breast, and other estrogen-responsive, neoplasms. In addition, fusion with CoRs should be an effective way of generating dominant negative mutants for other transcription factors or CoAs.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Plasmids
The reporter plasmid ERE2-tk109-luc has been described previously (90). GRE-tk109-luc was created by deleting the DR4 region from GRE-tk-DR4-luc, and rous sarcoma virus (RSV)-GR expression vector was used as previously described (91, 92). The pCMX-NCoR expression vector was provided by M. G. Rosenfeld (University of California, San Diego, CA) (33). pSG5-HEGO (wtER expression vector) was provided by Pierre Chambon (Université Louis Pasteur, Strasbourg, France) (93). The pCMX-NCoR-RD plasmid, containing the entire repressor domain region of NCoR, including RD1, RD2, and RD3 (a.a. 1–1551), was created by deleting a BstXI–SalI fragment from pCMX-NCoR and introducing an artificial EcoRI site. Fusion proteins of ER and NCoR were created as follows. To create pCMX-RD1–3-CDE, the CDE domains of ER (a.a. 181–553) were PCR-amplified from pSG5-HEGO by PCR using the primers, 5'-GATCGGTACCGCGGGCATGGAATTCGAGACTCGCTACTGTGCAGT-3' and 5'-CTAGCTAGGCGGCCGCTAGCGCTAGTGGGCGCATGTAGGC-3'. The PCR product was digested with EcoRI and NheI and inserted between the corresponding sites in pCMX-NCoR-RD. To create pCMX-RD1-CDE, repressor domain 1 (RD1) of NCoR (a.a. 1–312) was PCR amplified from pCMX-NCoR using the primers 5'-GATCGATCGCGGCCGCGCGGGCATGTCAAGTTCAGG-TTATCC-3' and 5'-CTAGGGATCCGCTAGCGAATTCATCATAACGTTGGCAGATTT-3'. Subsequently, the repressor domain region of pCMX-RD1–3-CDE was removed by restriction digestion and replaced with RD1.

pCMX-CDE-RD1 was constructed by ligating the KpnI–NotI fragment of CDE and the NheI–NotI fragment of RD1 into the pCMX expression vector. A DNA binding-deficient version of pCMX-CDE-RD1 was created by introducing three point mutations (E203G, G204S, and A207V) into the C region of CDE-RD1. pCMX-CDE was created by deleting the RD1 fragment of pCMX-CDE-RD1. pCMX-ER was constructed by cloning an EcoRI fragment of pSG5-HEGO, containing the entire cDNA for wtER, into pCMX. The dominant negative ER mutant L540Q was created by changing codon 537 of the ER gene from TAT (tyrosine) to TAG (stop). The codon for a.a. 540 was replaced by a stop codon to create the dominant negative ER mutant, ER1–536. pCMX-CAT was constructed by ligating the NotI fragment of pOP13CAT (Stratagene, La Jolla, CA), containing the cDNA for chloramphenicol acetyltransferase (CAT), into the NotI site of pCMX. The CAT gene product is unrelated to ER or NCoR and was used as a negative control. All constructs were verified by DNA sequencing.

Cell Culture
Sera and media were purchased from Life Technologies, Inc. (Gaithersburg, MD) and Sigma (St. Louis, MO). TSA-201 cells, derived from human embryonic kidney 293 cells (94) were grown in DMEM supplemented with 10% FBS. MDA-MB-231 (ER-negative) cells and T47D (ER-positive) cells, provided by V. Craig Jordan (Northwestern University Medical School, Chicago, IL), are human breast carcinoma cell lines. MDA-MB-231 cells were cultured in Eagle’s MEM supplemented with 5% calf serum, nonessential amino acids, and 10 mM HEPES. T47D cells were cultured in RPMI 1640 supplemented with nonessential amino acids and 10% FBS. Penicillin (100 U/ml) and streptomycin (100 µg/ml) were included in all media. Four days before transfection, cells were harvested using phenol red-free trypsin-EDTA and cultured in estrogen-depleted media (prepared without phenol red and supplemented with sera extracted three times with dextran-coated charcoal).

Transfections and Luciferase Assays
Cells were transferred to 12-well plates in estrogen-depleted medium 1 day before being transfected with ERE2-tk109-luc and expression vectors. Within each experiment, the total amount of DNA transfected into each group of cells was kept constant by the addition of the pCMX empty vector. TSA-201 cells were transfected with calcium phosphate and MDA-MB-231 cells with liposomes as previously described (90, 95). T47D cells were transfected by electroporation, using a Bio-Rad Laboratories, Inc. (Hercules, CA) Gene Pulser (300 V, 0.4-cm gap cuvette) with a capacitance extender (960 µfarads) and pulse controller (infinite resistance), and grown in estrogen-depleted medium for 24 h before treatment. E2 was added to treatment media as an ethanol stock solution, and ethanol was added to control wells to produce the same final solvent concentrations (typically 0.1%). Cells were treated for 40 (TSA-201) or 48 h (MDA-MB-231, T47D) and then assayed for luciferase activity as previously described (95).

Electrophoretic Mobility Shift Assays
Labeled probe was prepared by annealing two oligonucleotides (5'-CAAGTCAGGTCACAGTGACCTGATCAA-3' and 5'-TTGATCAGGTCACTGTGACCTGA-3') containing the Xenopus vitellogenin A2 ERE and filling the overhanging ends in the presence of [{alpha}-32P]dTTP (6000 Ci/mmol; Amersham Pharmacia Biotech, Arlington Heights, IL). Nuclear extracts were prepared as previously described (96) from TSA-201 cells. The cells were estrogen-depleted for 4 days, transfected with pCMX (no ER), pCMX-ER, pCMX-CDE, or pCMX-CDE-RD1 (20 µg DNA/15-cm plate) and incubated in estrogen-depleted medium for 2 days. Immediately before harvest, cells were treated for 2 h with 20 nM E2 to increase nuclear localization of the ER. Protein concentrations of the extracts were determined with a protein assay kit (Bio-Rad Laboratories, Inc.). Equal amounts (11 µg) of nuclear protein were used in each reaction. Extracts were preincubated with a binding buffer containing 20 mM HEPES, 40 mM KCl, 1 mM MgCl2, and 11 µg sperm DNA at 4 C for 30 min and then incubated with labeled ERE probe at 4 C for an additional 30 min, in a total volume of 20 µl. The monoclonal anti-ER antibody AER308 (NeoMarkers, Fremont, CA; Ref. 97) or a 100-fold excess of nonradioactive ERE probe was added in some of the experiments. The samples were loaded onto prerun 4% polyacrylamide gels, and radioactivity was visualized by autoradiography.

Western Blots and Immunocytochemistry
Western blots were performed on the same nuclear extracts described above. The extracts were fractionated in 4–15% SDS-PAGE ready gel (Bio-Rad Laboratories, Inc.) and transferred onto hydrophobic polyvinylidene difluoride membranes (Amersham Pharmacia Biotech). Immunodetection was performed using rat monoclonal ER antibody H222 and a LumiGLO kit (Kirkegaard & Perry Laboratories, Gaithersburg, MD). The antibody and protocols for immunoblotting and immunocytochemistry were kindly provided by Geoffrey L. Greene (University of Chicago, Chicago, IL). This antibody reacts with an epitope in the E domain of the ER (98). For immunocytochemical staining, TSA-201 cells were grown in estrogen-depleted media in 10-cm plates and transfected with 10 µg of pCMX, pCMX-ER, pCMX-CDE, or pCMX-CDE-RD1. Twenty-four hours after the transfection, cells were replated in Lab-Tek chambered glass slides (Nalge Nunc International, Naperville, IL) and treated with 20 nM E2 for another 24 h. The cells were washed with PBS and fixed with 4% paraformaldehyde. After being permeablized with methanol and acetone, cells were incubated with H222 or normal rat IgG in 10% goat serum for 60 min and stained using a Histostain-Plus kit (Zymed Laboratories, Inc. South San Francisco, CA) according to the manufacturer’s directions.

Estrogen Binding Assays
CDE-RD1 and wtER were synthesized in vitro from the corresponding plasmids using a TNT Coupled Reticulocyte Lysate System (Promega Corp., Madison, WI). Translation products were assayed for 125I-estradiol binding by the charcoal absorption method (99), as previously described (90).

GST Pull-Down Assays
ER and CDE-RD1 were translated in vitro as described above in the presence of [35S]methionine. The central region of SRC-1 (a.a. 661–855) was fused to GST in the expression vector pGEX2TK (Pharmacia Biotech, Piscataway, NJ) to create a GST-SRC-1 fusion protein. GST and GST-SRC-1 were transformed into Escherichia coli BL21(DES)/pLys, grown to an OD600 of 0.8 and incubated with 0.1 mM isopropyl-ß-D-thiogalactopyranoside overnight at 30 C. The bacteria were resuspended in NET buffer (150 mM NaCl, 5 mM EDTA, and 50 mM Tris, pH 7.4) with proteinase inhibitors and sonicated for 15 sec. After centrifugation, the supernatants were incubated with glutathione-agarose (Sigma) for 30 min at 4 C. The supernatant was removed and the agarose was washed five times with NET buffer plus proteinase inhibitors. Expression and purification of fusion proteins were confirmed by Coomassie blue staining. 35S-labeled wtER or CDE-RD1 was incubated for 30 min at 4 C with agarose-bound GST or GST-SRC-1 in the presence of 10 nM E2 or control vehicle. After washing five times with NET buffer plus proteinase inhibitors, bound proteins were recovered, resolved by 10% SDS-PAGE, and visualized by autoradiography.

Construction of Retroviral Vector, Transient Production of Retrovirus, and Transduction of T47D Cells
CDE-RD1 was inserted into the multiple cloning site of the retroviral vector pLXSN (CLONTECH Laboratories, Inc. Palo Alto, CA), and empty vector was used as a control. pLAPSN (CLONTECH Laboratories, Inc.), a vector with the same backbone but containing the alkaline phosphatase gene, was used to optimize transfection and transduction conditions. After optimization, alkaline phosphatase expression (assayed with Western Blue Stabilized Substrate for Alkaline Phosphatase; Promega Corp.) was detected in up to 50% of T47D cells. The Phoenix amphotropic packaging cell line was purchased from American Type Culture Collection (Manassas, VA) under the authorization of G. P. Nolan (Stanford University, Stanford, CA). Phoenix cells were grown in estrogen-depleted medium for 4 days and plated in 10-cm tissue culture plates 24 h before transfection. CDE-RD1-containing retroviral vector or control vector was transfected into Phoenix cells by the calcium-phosphate/chloroquine method following the protocol provided by G. P. Nolan. Packaging cells were provided with fresh medium 24 h after transfection and incubated at 32 C overnight. Culture supernatants from packaging cells containing retroviral particles were collected and centrifuged at 1500 rpm for 5 min to remove cell debris. T47D cells were estrogen depleted for 4 days and plated at a density of 5 x 105 cells per 10-cm plate 1 day before the transduction. Transduction was then performed by incubating T47D cells overnight in viral medium containing 4 mg/ml polybrene (Sigma). T47D cells were transduced twice to increase transduction efficiency. pLAPSN was used in parallel to monitor the transfection and transduction. A portion of the viral medium was used to transduce NIH3T3 cells (a gift from Richard Longnecker, Northwestern University, Chicago, IL), and the viral titer was determined as described in the protocol provided by CLONTECH Laboratories, Inc. The maximum titer used for the transduction was 105 colony-forming units (cfu)/ml.

Growth Assay
T47D cells transduced with control or CDE-RD1 retrovirus were seeded into four 96-well plates at a density of 3,000 cells per well on day 0 and placed in a 37 C incubator. The following day (day 1), cell density in one of the plates was measured by the tetrazolium reduction assay (Promega Corp., Madison, WI) as described previously (90). E2 (1 nM) or control vehicle was then added to the other cells. At 48-h intervals, a plate was taken for measurement of cell density and fresh media were applied to the remaining plates.


    ACKNOWLEDGMENTS
 
We thank M. G. Rosenfeld for the pCMX-NCoR expression vectors, Pierre Chambon for the pSG5-HEGO plasmids, Peter Kopp for GRE-tk109-luc reporter, Ron Evans for the RSV-GR expression vector, Craig Jordan for breast cancer cell lines, Richard Longnecker for the NIH3T3 cell line, and Geoffrey L. Greene for the antibody H222 and protocols used for immunoblotting and immunocytochemistry. We are also grateful to Wade Johnson for advice with gel shift assays; Eun-Jig Lee for help with immunocytochemical staining; Joanne McAndrews and Rachel Duan for helpful discussions; and Jeff Weiss for the comments about the paper.


    FOOTNOTES
 
Address requests for reprints to: J. Larry Jameson, M.D., Ph.D., Division of Endocrinology, Metabolism, and Molecular Medicine, Northwestern University Medical School, Tarry Building 15–709, 303 East Chicago Avenue, Chicago, Illinois 60611.

This work was supported in part by NIH Grant DK-42144 and by Department of Defense Grant USAMRDC B4337379.

Received for publication October 16, 1998. Revision received August 11, 1999. Accepted for publication September 9, 1999.


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