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Cell, Tumor, and Stem Cell Biology |
1 Center for Biotechnology and 2 Division of Medical Nutrition, Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden
Requests for reprints: Johan Hartman, Center for Biotechnology, Department of Biosciences and Nutrition, Karolinska Institutet, Novum, S-141 57 Huddinge, Sweden. Phone: 46-86089150; Fax: 46-87745538; E-mail: johan.hartman{at}biosci.ki.se.
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Abstract |
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and ERß. ER mediates the proliferative effect of estrogen in breast cancer cells, whereas ERß seems to be antiproliferative. We engineered ER-positive T47D breast cancer cells to express ERß in a Tet-Off–regulated manner. These cells were then injected orthotopically into severe combined immunodeficient mice, and the growth of the resulting tumors was compared with tumors resulting from injecting the parental T47D cells that do not express ERß. The presence of ERß resulted in a reduction in tumor growth. Comparison of the ERß-expressing and non-ERß–expressing tumors revealed that the expression of ERß caused a reduction in the number of intratumoral blood vessels and a decrease in expression of the proangiogenic factors vascular endothelial growth factor (VEGF) and platelet-derived growth factor ß (PDGFß). In cell culture, with the Tet-Off–regulated ERß-expressing cells, expression of ERß decreased expression of VEGF and PDGFß mRNA under normoxic as well as hypoxic conditions and reduced secreted VEGF and PDGFß proteins in cell culture medium. Transient transfection assays with 1,026 bp VEGF and 1,006 bp PDGFß promoter constructs revealed a repressive effect of ERß at the promoter level of these genes. Taken together, these data show that introduction of ERß into malignant cells inhibits their growth and prevents tumor expansion by inhibiting angiogenesis. (Cancer Res 2006; 66(23): 11207-13) |
Introduction |
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(ER)–positive tumors are treated with tamoxifen, which blocks the action of ER, or with aromatase inhibitors, which block the synthesis of E2 (2–4). These treatments are effective but patients inevitably develop hormone-resistant, invasive tumors. Because ERß is expressed in the normal breast, it is pertinent to ask whether this receptor might also be used as pharmacologic target in the treatment of breast cancer.
In breast cancer cells as well as in various mouse models, estrogens via ER stimulate proliferation and inhibit apoptosis (5, 6), whereas ERß opposes the proliferative effect of ER in vitro (7, 8). Several studies have reported that during tumor development in the breast epithelium, the expression of ER increases, whereas that of ERß decreases (9, 10). For a tumor to grow, increased proliferation must be accompanied by increased blood supply and this is achieved by angiogenesis and increased blood microvessel density (11–13). Angiogenesis is an essential process in normal tissue growth and repair and is regulated by multiple factors, including supply of nutrients, vascular endothelial growth factor (VEGF), platelet-derived growth factor ß (PDGFß), basic fibroblast growth factor (bFGF), and oxygen tension. Estrogens are important stimulators of angiogenesis in health as well as disease. In the uterine endometrium, estrogens via ER increase the transcription of VEGF (14, 15) and stimulate angiogenesis. Tamoxifen is antiangiogenic (16, 17).
The question addressed in the present study is whether an established breast cancer cell, which does not express ERß, can be influenced to behave more like a normal cell upon reintroduction of ERß. To answer this question, we engineered T47D breast cancer cells with a tetracycline-regulated ERß expression vector. We show here that both proliferation of the cancer cell and its ability to grow as a solid tumor in severe combined immunodeficient (SCID) mice were reduced upon expression of ERß.
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Materials and Methods |
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Experimental animals and xenograft model. Cells from one confluent 150-cm2 Falcon cell culture flask per mouse with T47D-ERß or normal T47D cells of the same clonal origin was diluted with 200 µL normal medium + 200 µL Matrigel (BD Falcon, San Jose, CA). The cell suspension was injected into the mammary fat pad of 5-week-old pathogen-free SCID/beige mice (Taconic, Ry) on day 0. E2 pellets, 0.72 mg/pellet (IRA, Sarasota, FL), were injected s.c. in the neck with pellet trochar (IRA). After 4, 8, 16, or 30 days, the mice were sacrificed and the tumor volume was measured with caliper according to the formula length x width x height. All tumors were fixed in 4% paraformaldehyde and stored in 75% ethanol. After this, tissue was paraffin-embedded and subsequently sliced into 4.5-µm sections according to standard protocol. Tumors of sufficient size at necropsy were divided into three similar parts for preparation of mRNA and protein as described below.
Western immunoblotting. Cells were plated and grown in 150-mm plates until 30% confluence was reached. At time point 0, the cells were incubated with E2 with or without tetracycline. Harvesting and extraction were done at different times (0-36 hours) according to standard protocol. SDS-PAGE was done as described (18). The following primary antibodies were used: VEGF (Santa Cruz Biotechnology, Santa Cruz, CA), PDGFß (BD Biosciences, San Jose, CA), Ki67 (DakoCytomation, Carpinteria, CA), CD31 (BD PharMingen, San Diego, CA), ß-actin (Sigma), and ERß (chicken antibody raised in our laboratory).
Immunohistochemistry. Antigen retrieval was done by microwave boiling in 0.01 mol/L citric acid (pH 6.0) or with Proteinase K solution for CD31 staining. Endogenous peroxidase activity was blocked with 0.5% hydrogen peroxide. Primary antibody was diluted 1:100 to 1:500 in 3% bovine serum albumin and 0.3% Triton X-100 (see above for antibody specifications) and incubated overnight at 4°C, followed by incubation with a biotinylated secondary antibody (1:200) in 0.3% Triton X-100 for 1 hour. Biotinylated secondary antibodies used were anti-mouse IgG and anti-rat IgG, both from Vector Laboratories (Burlingame, CA), incubated (1:200) in 0.3% Triton X-100 for 1 hour. Sections were finally incubated in the streptavidin-horseradish peroxidase ABC complex (Vector Laboratories) for 1 hour, stained in 3,3'-diaminobenzidine, and counterstained with Mayer hematoxylin (Sigma, Poole, United Kingdom) before dehydration through ethanol, and mounted in dibutyl phthalate xylene.
Measurement of microvessel density and counting of Ki67- and ERß-positive cells. Microvessel density was calculated by first identifying the areas of highest vascularization with x10 objective in the periphery of CD31 immunohistochemically stained tumor slides. Microvessel density was then determined by counting the number of vessels with x20 objective in two independent fields per tumor. Vessels to be counted were identified by one of the following criteria: CD31 (mouse)–positive vascular formations or vascular formations containing luminal erythrocytes. Average microvessel density and SD of each group (±ERß) was measured. Counting of ERß/Ki67–positive cells on immunohistochemically stained slides was done by first identifying areas with most intensive staining. The number of Ki67- and ERß-positive cells in two independent fields per tumor slide was counted with x20 objective. Average number of stained cells and SD for each group (±ERß) were calculated.
ELISA assay. ERß-inducible T47D cells were incubated in estrogen-depleted RPMI medium containing 50 nmol/L ICI for 48 hours. Thereafter, medium was changed to normal RPMI medium ± E2, ±tetracycline, and cell cultures were placed in hypoxia chamber containing <1% O2 at 37°C. After 72 hours, the conditioned cell medium was analyzed according to standard ELISA protocol (Quantakine human VEGF and PDGFß; R&D Systems, Minneapolis, MN). RNA was extracted and purified from the T47D cells according to standard protocol.
Real-time PCR. RNA extraction and cDNA synthesis were done as described earlier (19). Real-time PCR was done with SYBR-Green PCR Master Mix or TaqMan Universal Mastermix (Applied Biosystems, Foster City, CA). The following primers and probes were used: VEGF forward: 5'-CTCTACCTCCACCATGCCAAGT-3'; reverse: 5'-TGATTCTGCCCTCCTCCTTCT-3'. PDGFß forward: 5'-CTGCTACCTGCGTCTGGTCA-3'; reverse: 5'-CATCAAAGGAGCGGATCGA-3. ERß forward: 5'-TCCATGCGCCTGGCTAAC-3'; reverse: 5'-CAGATGTTCCATGCCCTTGTTA-3. Probe: 5'-FAM (6-carboxyfluorescein)-TCCTGATGCTCCTGTCCCACGTCA(6-carboxytetramethylrhodamine)-3'. 18S rRNA forward: 5'-CCTGCGGCTTAATTTGACTCA-3'; reverse: 5'-AGCTATCAATCTGTCAATCCTGTCC-3 as a reference gene. The real-time PCR reactions were done in an ABI PRISM 7500 (Applied Biosystems) under the following conditions: 50°C for 2 minutes, 95°C for 10 minutes, followed by 40 to 50 cycles at 95°C for 15 seconds and 60°C for 1 minute. The optimum concentration of primers was determined in preliminary experiments and all primer pairs were checked with melting curve analysis.
Cloning and transfections. The VEGF promoter (forward primer, 5'-AAATTCTTCTCCCCTGGGAA-3'; reverse primer, 5'-AATGAATATCAAATTCCAGCA-3') and PDGFß promoter (forward primer, 5'-CAGTGCAAGCGGAGGAGATGA-3'; reverse primer, 5'-CGGCTGCAGGAGGAGAAGTTG-3') were amplified by PCR from human genomic DNA and subsequently subcloned into pGL3-basic luciferase vector (Promega).
Transfection of T47D cells was done by culturing cells on six-well cell culture dishes until 50% to 70% confluence was reached, whereupon the cells were synchronized. Transient transfections were done using Lipofectamine 2000 (Invitrogen) according to the protocol from the manufacturer. The reporter plasmid (VEGF or PDGFß promoter 1.0 µg/well) was transiently cotransfected together with an expression plasmid (ERß 0.5-0.1 µg) for 6 hours. When necessary, the total amount of DNA was completed with pcDNA3 (Invitrogen). After transfection, cells were treated with 10 nmol/L E2 or ethanol in stripped medium for 24 or 48 hours. Cells were then lysed and luciferase activity was determined with a microplate luminometer (Berthold) using luciferase assay kit from Biothema. All data were normalized to ß-galactosidase enzyme activity obtained after cotransfection with a ß-galactosidase-lacZ plasmid (100 ng/well).
Hypoxia chamber experiments. T47D-ERß cells were incubated for 48 hours in normal stripped medium without E2 during normoxia. Cells were then incubated in 0.5% O2 in 37°C for 48 hours in the presence of E2. Cells were subsequently harvested with TRIzol reagent according to instructions from the manufacturer.
Statistics. Values are expressed as means with 95% confidence intervals. Tumor growth curves were constructed from the mean tumor volume at each time point of measurement. Unpaired, two-tailed t test was used to compare differences between two groups. One-way ANOVA analysis with Tukey's multiple comparison posttest was used to test differences between three or more groups. Significance is presented as *P < 0.05; **P < 0.005; ***P < 0.001; and NS, nonsignificant.
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Results |
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-positive breast cancer cells such as T47D or MCF7, E2 treatment leads to increased proliferation and increased tumor growth when these cells are implanted into mice (20, 21). To study the effect of ERß in this context, we have generated T47D breast cancer cells with stable Tet-Off–inducible ERß expression. These cells are grown in the presence of tetracycline and should only express high levels of ERß after removal of tetracycline. By orthotopic implantation of T47D-ERß or parental T47D cells into SCID/beige mice, the effect of ERß on ER-induced tumor growth may be investigated. Preliminary studies with T47D-ERß cells implanted into SCID/beige mice showed that there was some expression of ERß even in the presence of tetracycline. Accordingly, tumor growth in the presence of tetracycline could not be used as baseline measure of ER-controlled growth and parental T47D cells were used instead. T47D cells are E2 dependent for their growth; therefore, at the start of the experiment, all animals were implanted with 60-day-release E2 pellets. To confirm that ERß was expressed in T47D-ERß tumors, ERß mRNA and protein levels were monitored by real-time PCR and immunohistochemical analysis. There was an extremely low, but nonetheless measurable, level of ERß mRNA in T47D cells. As shown in Fig. 1A , exogenous ERß expression reduced tumor volume at end point 30 days by 80%. Of the eight mice implanted with T47D-ERß, in three animals at necropsy, no tumor tissue could be found. Thirty days after implantation, levels of ERß mRNA in T47D-ERß xenographs were >10-fold higher than in the T47D xenografts (Fig. 1B). Immunohistochemical detection of ERß in paraffin-embedded tumor slides showed strong expression of ERß 4 to 16 days after implantation. There was a gradual decline in expression so that by day 30, there was only a small but statistically significant difference in ERß expression between the T47D-ERß and the T47D tumors (P = 0.0421; Fig. 1C).
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ERß decreases expression of VEGF and PDGFß in T47D-ERß cells. We investigated whether ERß was directly involved in regulating the production of VEGF and PDGFß or whether the induction of these factors was induced by hypoxia independent of the presence of ERß. We incubated ERß-inducible T47D cells in vitro under normoxic conditions (20% O2) in presence of E2. Expression of ERß reduced VEGF mRNA as well as in secreted form under both normoxic and hypoxic conditions (Fig. 4A and B
). PDGFß was strongly down-regulated at the mRNA level and at secreted protein level as a result of ERß expression, both under normoxic and hypoxic conditions (Fig. 4C and D). To further investigate this transcriptional regulation, cells were transiently transfected with promoter constructs of VEGF and PDGFß. E2 treatment resulted in an increase in promoter activity with both constructs (Fig. 5
), caused by ligand activation of ER. Cotransfection of an ERß expression vector resulted in reduction of VEGF and PDGFß promoter activities in the presence of E2.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-mediated cell cycle progression by interference with cell cycle factors such as cyclin E and Cdk2 (7). The purpose of the present study was to determine whether the observations made in cell culture would apply to breast cancer cells in vivo. As shown by others using viral transfection systems, ERß inhibits tumor formation of MCF-7 cells in a mouse model (8). One critique of viral transfection studies is that uncontrolled overexpression of the transfected gene leads to unphysiologically high protein levels. With the use of a Tet-Off-system, we here report that when ERß expression is induced in breast cancer xenografts, tumor growth is inhibited. Furthermore, the proliferation marker Ki67 was inversely associated with ERß in most groups; when the number of ERß+ cells was high, the number of Ki67+ cells was low, whereas when the number of ERß+ cells was low, the number of Ki67+ cells was high (Fig. 1C and D). This finding gives further support to the notion that ERß is antiproliferative. There was a decrease in the number of ERß+ tumor cells from days 16 to 30 in the T47D-ERß group (Fig. 1C), in spite of strong induction of ERß at mRNA level. The reason for this loss is not known but a possible mechanism could be increased ERß protein degradation. This would also be in line with the reports showing decreased ERß levels in breast cancer. Studies of the mechanisms behind the observed decrease in ERß levels in the transplanted tumor cells are ongoing.
One possible explanation for the reduction in tumor volume by ERß expression could be the influence of ERß on angiogenesis. We found that in the T47D-ERß xenografts, microvessel density was significantly lower than in the parental T47D tumors (Fig. 3A). As expected, PDGFß was reduced on both mRNA and protein level in the T47D-ERß xenografts. We could see no reduction of VEGF-mRNA in xenografts by ERß expression but a nonsignificant trend toward a reduction of VEGF protein (P = 0.0723; Fig. 3B and C).
Tamoxifen, which is known to activate ERß in certain promoter contexts, has been shown to inhibit growth of breast cancer xenografts by affecting angiogenesis (25). Because oxygen tension is a major stimulator of angiogenesis, the question was whether the lower microvessel density in the ERß-expressing tumors was simply because the tumors were smaller and less hypoxic. We therefore analyzed the expression of the three most important proangiogenic factors, namely VEGF, bFGF, and PDGFß, during normoxic as well as hypoxic conditions in vitro. Secreted bFGF protein was not detectable in the cell culture supernatants but both PDGFß and VEGF were reduced by ERß expression (Fig. 4).
It has earlier been reported that ER increases the expression of PDGFß, bFGF, and VEGF, and that this induction of growth factors might be an important mechanism explaining how ER stimulates breast cancer growth and progression. The role of ERß in regulation of PDGFß is not clear; furthermore, there are conflicting data as to how ERß affects VEGF expression in tumors (26, 27).
In transient transfections with a 1,026 bp VEGF promoter construct, a reduction in promoter activity by cotransfection with ERß was seen in the presence of E2. Accordingly, ERß might be a direct transcriptional inhibitor of VEGF, opposing the effect of ER. E2 treatment in the absence of ERß cotransfection showed only a minor up-regulation of the VEGF promoter activity. Earlier reports with longer promoter constructs have shown strong activation by ER (28). VEGF is well known for its activation of endothelial cells, resulting in their proliferation and migration. In addition, as shown in a recent report (29), VEGF influences breast cancer cell survival and growth by direct effects on cancer cell surface VEGFR-2 in an autocrine fashion.
Transient transfections with 1,006 bp PDGFß promoter constructs also showed inhibition by ERß. PDGFß is a growth factor with wide potentials. In addition to its proangiogenic effects, PDGF family members stimulate proliferation of breast cancer cells (30). Also, PDGFß have been reported to induce aromatase activity in T47D cells (31). Therefore, it is an interesting possibility that induction of ERß would actually inhibit the local production of E2 in breast tumors, in the same way as pharmaceutical aromatase inhibitors do. Reduction of growth factor expression could provide one explanation as to how ERß inhibits tumor growth. Furthermore, preliminary experiments done by our group indicate that ERß correlates inversely to PDGFß and VEGF mRNA in purified breast cancer cells from 14 different human breast tumors. This will be investigated in more detail with increased sample size.
Altogether, our results indicate an antitumorigenic role of ERß in breast cancer. This makes ERß an interesting therapeutic target in breast cancer and perhaps treatment with ERß-selective ligands might work as both antiproliferative and antiangiogenic therapy.
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Acknowledgments |
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The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
We thank Margaret Warner for her advice and critical comments on the manuscript.
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Footnotes |
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Received 1/ 4/06. Revised 9/17/06. Accepted 10/ 3/06.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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activation function-1 induces proliferation of breast cancer cells. J Biol Chem 2003;278:26704–14. (ER) and ß (ERß) differentially regulate proliferation and apoptosis of the normal murine mammary epithelial cell line HC11. Oncogene 2005;24:6605–16.[CrossRef][Medline] and ß. Cancer Res 2002;62:4977–84. and hypoxia-inducible factor 1 to the vascular dothelial growth factor promoter. Mol Endocrinol 2005;19:2006–19.This article has been cited by other articles:
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  M. Dadiani, D. Seger, T. Kreizman, D. Badikhi, R. Margalit, R. Eilam, and H. Degani Estrogen regulation of vascular endothelial growth factor in breast cancer in vitro and in vivo: the role of estrogen receptor {alpha} and c-Myc Endocr. Relat. Cancer, September 1, 2009; 16(3): 819 - 834. [Abstract] [Full Text] [PDF]
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 J. Hartman, K. Edvardsson, K. Lindberg, C. Zhao, C. Williams, A. Strom, and J.-A. Gustafsson Tumor Repressive Functions of Estrogen Receptor {beta} in SW480 Colon Cancer Cells Cancer Res., August 1, 2009; 69(15): 6100 - 6106. [Abstract] [Full Text] [PDF]
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 A. M. Sotoca Covaleda, H. van den Berg, J. Vervoort, P. van der Saag, A. Strom, J.-A. Gustafsson, I. Rietjens, and A. J. Murk Influence of Cellular ER{alpha}/ER{beta} Ratio on the ER{alpha}-Agonist Induced Proliferation of Human T47D Breast Cancer Cells Toxicol. Sci., October 1, 2008; 105(2): 303 - 311. [Abstract] [Full Text] [PDF]
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 N. Honma, R. Horii, T. Iwase, S. Saji, M. Younes, K. Takubo, M. Matsuura, Y. Ito, F. Akiyama, and G. Sakamoto Clinical Importance of Estrogen Receptor-{beta} Evaluation in Breast Cancer Patients Treated With Adjuvant Tamoxifen Therapy J. Clin. Oncol., August 1, 2008; 26(22): 3727 - 3734. [Abstract] [Full Text] [PDF]
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 C. M. Klinge, N. S. Wickramasinghe, M. M. Ivanova, and S. M. Dougherty Resveratrol stimulates nitric oxide production by increasing estrogen receptor {alpha}-Src-caveolin-1 interaction and phosphorylation in human umbilical vein endothelial cells FASEB J, July 1, 2008; 22(7): 2185 - 2197. [Abstract] [Full Text] [PDF]
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 D. J. Kroll, H. S. Shaw, and N. H. Oberlies Milk Thistle Nomenclature: Why It Matters in Cancer Research and Pharmacokinetic Studies Integr Cancer Ther, June 1, 2007; 6(2): 110 - 119. [Abstract] [PDF]
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