Estrogen-receptor-dependent regulation of telomerase activity in human endometrial cancer cell lines

Gynecologic Oncology 103 (2006) 417 – 424 www.elsevier.com/locate/ygyno

Estrogen-receptor-dependent regulation of telomerase activity in human endometrial cancer cell lines☆ John F. Boggess a,⁎, Chunxiao Zhou a,1 , Victoria L. Bae-Jump a , Paola A. Gehrig a , Young E. Whang b a

Department of Obstetrics and Gynecology, Division of Gynecologic Oncology, Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, CB# 7570, Chapel Hill, NC 27599, USA b Department of Medicine, Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA Received 16 November 2005 Available online 11 May 2006

Abstract Objectives. Given that prolonged exposure to unopposed estrogen is associated with endometrial cancer development and that the promoter region of the catalytic subunit of the telomerase enzyme, hTERT, contains putative estrogen response elements (EREs), we postulated that estrogen-receptor (ER)-mediated induction of telomerase activity may play an important role in endometrial carcinogenesis. Methods. ER-positive and ER-negative endometrial cancer cell lines were used. ERα expression was reconstituted in ER-negative cell lines by transient transfection. Telomerase activity was assayed using a PCR-based telomeric repeat amplification protocol (TRAP) after exposure to estradiol (E2). hTERT mRNA expression was assessed by real-time RT-PCR. Gel shift assays using oligonucleotide probes encoding each ERE and transient expression assays using luciferase reporter plasmids containing varying lengths of the 5′ promoter region of the hTERT gene were performed. Results. E2 induced both hTERT gene transcription and telomerase activity in the ER-positive cell lines, but not in the ER-negative cell lines. Transfection of ERα into ER-negative cell lines restored E2-induced hTERT gene transcription and telomerase activity. Gel shift assays revealed two EREs in the hTERT promoter that specifically bind to ERα. Luciferase assays demonstrated that at least the proximal ERE is responsible for transcriptional activation by ligand-stimulated ERα. Conclusions. Telomerase activity and hTERT mRNA were increased in response to estrogen in an ERα-dependent fashion in endometrial cancer cells. Binding of complexed estrogen with ERα to the EREs found within the hTERT promoter suggests a possible mechanism for telomerase induction that may facilitate the malignant transformation of hormone-dependent endometrial cells. © 2006 Elsevier Inc. All rights reserved. Keywords: Endometrial cancer; Telomerase; Estrogen; Estrogen receptor

Introduction Telomeres are specialized nucleoprotein structures that cap the ends of eukaryotic chromosomes. Due to the inability of DNA polymerase to replicate the ends of double-stranded DNA, ☆

Supported by NIH Grant 5K08CA085772 awarded to Young E. Whang and V Foundation Grant awarded to Paola A. Gehrig. ⁎ Corresponding author. Fax: +1 919 966 5670. E-mail addresses: [email protected] (J.F. Boggess), [email protected] (V.L. Bae-Jump). 1 This author contributed equal work to manuscript as first author. 0090-8258/$ - see front matter © 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ygyno.2006.03.032

telomeres progressively shorten with each cell division. This process ultimately leads to cell senescence and apoptotic cell death. Telomerase is a ribonucleoprotein enzyme that compensates for telomere shortening during cell division by synthesizing telomere DNA and thus maintaining telomere length. In most normal somatic cell types, telomerase activity is usually undetectable [1]. Examples of the limited number of benign somatic tissues that express telomerase include the epidermis, bone marrow and endometrium. It is thought that telomerase plays a critical role in the ability of normal endometrium to repeatedly proliferate from the onset of menarche to menopause. Furthermore, activation of telomerase has also been implicated

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in many cancers, including gynecologic malignancies [2,3], as a fundamental step in cellular immortality and oncogenesis [1]. The human endometrium is composed of epithelial glands and connective tissue, which demonstrate complex patterns of proliferation, secretory activity and breakdown in response to estrogen and progesterone exposure and withdrawal. Telomerase is expressed by the endometrial epithelial glands differentially throughout the menstrual cycle with high levels of expression observed in the proliferative phase when serum estrogen levels are maximal followed by near absent levels by the middle of the secretory phase when progesterone levels are high [4]. Observational studies have suggested that estrogen activates and progesterone suppresses telomerase activity in normal endometrium [5–7]. Telomerase is comprised of an RNA template (hTR) and the catalytic protein hTERT which has reverse transcriptase activity. hTERT is considered to be the most important factor in the formation of functional telomerase [8]. Transcription of hTERT is regulated by many transcription factors such as p53, E2F and steroid hormone receptors [8]. It has been previously demonstrated that estrogen can activate telomerase through the direct interaction of the hormone-activated estrogen receptor (ER) with estrogen receptor elements (EREs) in the hTERT 5′ regulatory region in ER-positive breast cancer cells, ovarian cancer cells and human ovarian epithelium [9–11]. Endometrial cancer is the most common gynecologic malignancy diagnosed in the United States. It is estimated that 40,880 new cases will be diagnosed in 2005, and 7310 women will succumb to this disease [12]. Endogenous and exogenous estrogen exposure are major risk factors for the development of endometrial cancer [13]; however, the molecular link between estrogen and endometrial carcinogenesis remains poorly understood. Given that prolonged exposure to unopposed estrogen is associated with excess proliferation and ultimately the development of endometrial cancer and that the promoter region of the catalytic subunit of the telomerase enzyme, hTERT, contains putative EREs [9,10], we postulate that ER-mediated induction of telomerase activity may play an important role in the pathogenesis of endometrial cancer. The aim of this study was to elicit the underlying mechanisms involved in ER regulation of hTERT transcription and telomerase activity in ER-positive and ERnegative endometrial cancer cell lines. Materials and methods Cell culture The regulation of telomerase expression was investigated in ER-positive (Ishikawa, ECC-1) and ER-negative (RL 95-2, HEC-1B) endometrial cancer cell lines. All of these cell lines were provided by Dr. Bruce Lessey (Center for Women's Medicine, Greenville, SC). As previously described, estrogen-induced ERE chloramphenicol acetyltransferase (CAT) activity was determined in each of these cell lines to confirm functional ER status [14]. RL 95-2 cells were grown in a 1:1 mixture of Dulbecco's minimal essential medium and Ham's F12, supplemented with 10% fetal bovine serum (FBS), 5 μg/ml of bovine insulin, 100 units/ml penicillin and 100 μg/ml streptomycin, in the presence of 5% CO2 at 37°C. Ishikawa and HEC-1B cells were grown in EMEM supplemented with 5% FBS, 5 μg/ml of bovine insulin, 100 units/ml penicillin and 100 μg/ml streptomycin in the presence 5% CO2 at 37°C. ECC-1 cells were

grown in RPMI 1640 medium containing 5% FBS and 6 mM NaCO3 in the presence 5% CO2 at 37°C. All human endometrial cancer cell lines were cultured in phenol-red free medium with 0.5% charcoal-dextran-treated FBS for 1 day before treatment with estrogen.

Chemicals and plasmid Estrogen receptor alpha (ERα) expression vector was obtained from Dr. Carolyn Smith (Baylor College of Medicine, Houston, TX). All hTERT reporter promoter luciferase plasmids were provided by Dr. I. Horikawa (National Institute of Health, Bethesda, MD). 32P-labeled ATP (3.000 ci/mmol) was obtained from Amersham Pharmacia Biotech (Arlington Heights, IL). 17-β estradiol (E2) was purchased from Sigma (St. Louis, MO). All other chemicals were from Sigma (St. Louis, MO).

Estrogen treatment Cells were seeded at 4 × 105 cells per T25 culture flask or 1.5 × 105 cells per well of a 12-well plate, containing either 5 ml or 1.2 ml regular medium, respectively, and then changed to phenol-red free medium with 0.5% stripped FBS for incubation at 37°C overnight. Immediately prior to treatment, the medium in the culture plates was aspirated, triply washed with phosphatebuffered saline (PBS) and replaced with fresh medium. E2 dissolved in ethyl alcohol was added to each well at concentrations ranging from 0.01 to 10 μM and incubated for 6–96 h. Concurrently, the same amount of ethyl alcohol was added to the control wells.

Estrogen receptor transfection Estrogen receptor transient transfection was performed by the calcium phosphate method with mammalian transfection systems kits from Promega Corporation (Madison, WI). Briefly, on the day before transfection, RL 95-2 or HEC-1B cells were plated at a density of 1 × 104/cm2 in 100 mm plates. Three hours prior to the addition of vector, fresh medium containing 10% FBS was provided to the cells. Calcium phosphate–DNA precipitates were prepared as indicated in the protocol. Precipitates were allowed to stand for 30 min without agitation before being added to the cell plates.

Telomerase activity assay Telomerase activity in cultured cells was assayed using a PCR-based telomeric repeat amplification protocol (TRAP) as described previously [4]. The TRAP-eze Telomerase Detection Kit (Intergen, Purchase, NY) was used as recommended by the manufacturer with minor modifications. Briefly, 1 × 105 cell pellets were stored at − 80°C until lysis was performed. Samples were homogenized with 1× CHAPS lysis buffer (3-[(3-cholamidopropyl)dimethylammonio]-1-propane-sulfonate) at 1 × 105 cells/100 μl, incubated on ice for 30 min and centrifuged at 13,000×g for 21 min at 4°C. The supernatant was transferred into a fresh tube, and the protein concentration was measured by ABC kit (Bio-Rad, Hercules, CA). Between 0.25 and 0.5 μg of protein from the cell extract was mixed with 32P-labeled TS primer in a 50 μl reaction mixture. After 30 min of incubation at 30°C, PCR amplification was then performed with 27 cycles at 94°C for 30 s and 59°C for 30 s. The PCR products were separated by electrophoresis on 10% polyacrylamide non-denaturing gels. Phosphor-Imager and Imagequant software from Molecular Dynamics (Sunnyvale, CA) were used to quantify the band intensities. Telomerase activity is expressed quantitatively as TPG, which reflects a ratio of the TRAP product ladder bands to the internal telomerase assay standard band and was calculated according to the formula supplied in the manufacturer's manual. The reliability and linearity of TPG as a measure of telomerase activity have been confirmed by other investigators [15].

Real-time RT-PCR for hTERT Total RNA was isolated using the RNAqueous kit (Ambion, Austin, TX) and further purified by the DNA-free kit (Ambion, Austin, TX). The reverse transcription and PCR reactions were performed using the TaqMan Gold onestep RT-PCR kit in the ABI Prism 7700 Sequence Detection System (Applied

J.F. Boggess et al. / Gynecologic Oncology 103 (2006) 417–424 Biosystems, Foster City, CA). Reverse transcription was carried out at 48°C for 30 min. The PCR condition consisted of a 10-min step at 95°C and 40 cycles at 95°C for 15 s and 65°C for 1 min. A housekeeping control gene acidic ribosomal phosphoprotein P0 (RPLP0, also known as 36B4) was used as an internal control to correct for differences in the amount of RNA in each sample [16]. Primers and fluorogenic probes for hTERT and RPLP0 have been described previously [16]. The standard curve for hTERT was generated by using dilutions of a known amount of cRNA synthesized by in vitro transcription of a cloned fragment. The normalized level of hTERT in each sample was estimated by a ratio of the hTERT level to the RPLP0 level, as described previously [16].

Gel shift assay Electrophoretic mobility shift assays were performed using the Gel Shift Assay System (Promega, Madison, WI) according to the manufacturer's instructions with minor modifications. Ten micrograms of nuclear protein extracts from the Ishikawa and ERα-transfected RL 95-2 cells was incubated with 17.5 fmol 32P-labeled double-stranded oligonucleotides containing either an ERE in the core promoter at − 2777/− 2755 (5′-TGTTGGTCAGGCTGATCTCAAA) or an ERE/Sp1 at − 979/−956 (5′-ATCTGGTCACATCCCGCCCGCACA) [9,10]. Specificity of binding was determined by adding excess unlabeled oligonucleotides or the non-specific competitor Ap2. DNA–protein binding complexes were analyzed on 6% polyacrylamide gels. The gels were dried and subjected to autoradiography on either X-ray films or a Phosphor-Imager screen.

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Luciferase assay Transient transfection of luciferase reporter plasmids was performed using the TransFast Transfection Reagent (Promega, Madison, WI). The Ishikawa and ERα-transfected RL 95-2 cells were used in these experiments. Briefly, 8 × 104 cells were seeded in 24-well plate overnight and transfected with promoter luciferase plasmids (0.5 μg/well). The pRL-SV40 (2 ng/well) containing Renilla reniformis luciferase was co-transfected in each transfection as an internal control to normalize the transcriptional activity of the hTERT promoter plasmids. The luciferase assay was then performed using the Dual Luciferase Reporter Assay System (Promega, Madison, WI) according to protocols provided by the manufacturer. All experiments were performed at least three times for each plasmid.

Results Effect of estrogen on telomerase activity and hTERT expression in endometrial carcinoma cell lines E2 exposure resulted in increased telomerase activity by TRAP assay in our ER-positive endometrial cancer cell lines (Ishikawa and ECC-1). The increased activity was dependent on

Fig. 1. (A) Effect of estrogen on telomerase activity among ER-positive (Ishikawa and ECC-1) and ER-negative (RL 95-2 and HEC-1B) endometrial cancer cell lines. Cells were treated with different concentrations of E2 for 48 h, and telomerase activity was assessed by TRAP assay. (B) Above data represented in graphical form using TPG (total product generated) which corresponds to relative telomerase activity. TPG is calculated from the ratio of TRAP product band to the internal telomerase assay standard band (ITAS). Data shown represent at least two independent experiments.

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Fig. 2. Effects of estrogen on telomerase activity in ER-positive Ishikawa and ECC-1 cells. (A) Cells were treated with E2 (1 μM) for 6–72 h in a time course fashion. Telomerase activity was determined by the TRAP assay. (B) Relative telomerase activity was quantitated using TPG in the Ishikawa and ECC-1 cells treated with 0.1, 1 and 10 μM E2 for 6–72 h. (C) hTERT mRNA expression after E2 exposure at different concentrations was assessed by real-time RT-PCR in the ER-positive Ishikawa and ECC-1 cells. Data are presented as means ± standard deviation (SD) of duplicated samples from at least two independent experiments.

both E2 concentration and time of exposure (Figs. 1, 2A and B). Maximum telomerase activity was detected at E2 concentrations ranging from 0.1 to 10 μM at 48 h after treatment. The increased

activity was detected first at 12 h, appeared to peak after 48 h and remained elevated up to 72 h after exposure to estrogen. No effect was detected when E2 was present for only 6 h or at

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Fig. 3. Telomerase activity and hTERT expression in estrogen-stimulated ERα-transfected RL 95-2 cells. Cells were treated with E2 at 0.01 to 10 μM for 48 h. (A) Left panel, TRAP assay products separated by gel electrophoresis. Right panel, relative telomerase activity (TPG). C1, non-transfected control. ERα-transfected RL 95-2 cells were assessed by TRAP assay (B) or by real-time RT-PCR (C) after exposure to varying concentrations of E2. Data are presented as means ± standard deviation (SD) of duplicated samples from at least two independent experiments.

concentrations less than 0.1 μM. No effect of E2 on telomerase activity by TRAP assay was detected in the ER-negative endometrial cancer cell lines (RL 95-2 and HEC-1B) (Fig. 1). To understand the mechanisms underlying loss of telomerase activity, we quantified by real-time RT-PCR the level of hTERT mRNA. The hTERT gene encodes the catalytic subunit of telomerase and is usually the rate-limiting determinant of telomerase enzymatic activity. Treatment with E2 induced the hTERT mRNA level in a dose-dependent manner in the ERpositive endometrial cancer cell lines (Ishikawa and ECC-1) (Fig. 2C). Maximum up-regulation of hTERT was seen at E2 concentrations ranging from 0.1 to 10 μM at 48 h after treatment. An increase in hTERT mRNA was seen as early as 12 h after exposure to E2. hTERT mRNA levels peaked 48 h after exposure to E2 and were persistently elevated up to 72 h. No effect was detected when E2 was present for only 6 h or at concentrations less than 0.1 μM. E2 did not increase hTERT mRNA expression in the ER-negative endometrial cancer cell lines (RL 95-2, HEC-1B) (data not shown). In order to further investigate the role of the ER in estrogen's apparent effect on telomerase activity in endometrial cancer cell

lines, the ER-negative RL 95-2 cells were transfected with ERα. Following transfection with ERα, E2 increased telomerase activity as quantified by the TRAP assay in a dose-dependent and time-dependent fashion similar to the ER-positive endometrial cancer cell lines (Ishikawa and ECC-1). Within 48 h of exposure to 0.1 μM of E2, a six-fold increase in telomerase activity was evident in the ERα-transfected RL 95-2 cells compared to the uninfected RL 95-2 cells (Figs. 3A and B). Increased hTERT mRNA expression was also seen in response to E2 in the ERα-transfected RL 95-2 cells but not in the uninfected control cells (Fig. 3C). These results demonstrate that ERα is necessary and sufficient to confer E2-dependent regulation of hTERT gene transcription in endometrial cancer cell lines. Estrogen receptor binds proximal and distal ERE sequences in the hTERT promoter The regulatory promoter sequence of the hTERT gene contains two putative EREs in the 5′ flanking sequence: the distal one at − 2777/− 2755 and the proximal one at − 979/− 956

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Fig. 4. Identification of two EREs in the hTERT promoter by gel shift assay. Nuclear extracts from the ER-positive Ishikawa (A, B) or ERα-transfected RL 95-2 cells were subjected to gel shift using two EREs as probes. (A) Gel shift using 32P-labeled double-stranded oligonucleotide containing an ERE at − 2777/− 2755 in Ishikawa cells. Lane 1, 1 μg protein. Lane 2, 3, 1 μg protein + 50 and 5-fold molar excess of unlabeled probe. Lane 4, 1 μg protein + Ap2 (non-specific competitor). (B) Using oligonucleotide probe containing half ERE at − 979/− 956 in Ishikawa cells. Lane 1, 1 μg protein. Lane 2, 3, 1 μg protein + 50 and 5-fold molar excess of unlabeled probe. Lane 4, 5 μg protein + Ap2. (C) Using oligonucleotide probe containing entire ERE in ERα-transfected RL 95-2 cells. Lane 1, 1 μg protein from non-transfected RL 95-2 cells. Lane 2, 1 μg protein from ERα-transfected RL 95-2 cells. Lane 3, 1 μg protein + 50-fold molar excess of unlabeled probe. Lane 4, 1 μg protein + Ap2. Lane 5 is negative control for each. Data are presented as means ± standard deviation (SD) of duplicated samples from at least two independent experiments.

that overlaps with an Sp1 binding site [9,10]. To determine whether ERα binds to these sites, gel shift analysis was performed. Gel shift assays performed with an oligonucleotide probe encoding each ERE and nuclear extracts from the ERpositive Ishikawa cells or the ERα-transfected RL 95-2 cells demonstrated that DNA binding proteins, likely ER and associated proteins, are capable of binding to both the proximal and distal ERE sequences (Fig. 4). A specific band was observed with probes containing the putative ERE at − 2777/− 2755 or the ERE/Sp1 site at − 979/− 956, both of which were competed by increasing concentrations of unlabeled probe but not by the nonspecific competitor Ap2. These findings suggest that ERα regulates telomerase activity in endometrial cancer cells by binding directly to the hTERT promoter through each of these EREs. Functional characterization of the hTERT EREs In order to delineate the functional significance of the ERE sequences, we performed a series of transient transfections with luciferase reporter plasmids containing the varying lengths of the 5′ promoter region of the hTERT gene (Fig. 5). When the ERpositive endometrial cancer cells (Ishikawa and ECC-1) were transfected with reporter plasmids and stimulated with estrogen, reporter constructs containing no promoter or the 5′ sequence without either ERE (i.e. P88, P255 and P385) expressed low basal luciferase activity and were unresponsive to E2 treatment. In contrast, E2 stimulated luciferase activity from larger reporter constructs containing the proximal ERE (i.e. P1125, P3195 and P3328) by about 4- to 5-fold. These data imply that the proximal

ERE is sufficient to confer E2 responsiveness to hTERT transcription in endometrial cancer cells. Similar results were also seen in the ERα-transfected RL 95-2 cells. However, we cannot rule out the role that the distal ERE plays in this process. Discussion In this work, we provide evidence that estrogen directly regulates telomerase activity and hTERT gene expression in a series of ER-positive and ER-negative endometrial cancer cell lines. To our knowledge, this is the first study to confirm this relationship in endometrial cancer cells. In the ER-positive Ishikawa and ECC-1 cells, E2 stimulated telomerase activity and increased hTERT mRNA levels. These findings confirm those in other recent studies that have reported that estrogen activates telomerase in breast cancer cells [10], ovarian cancer cells [11] and human ovarian epithelium [9]. Given that telomerase activity and hTERT mRNA levels rose in parallel, this suggests that estrogen stimulates telomerase by regulation of transcription of the hTERT gene. In contrast, no significant increase in telomerase activity or hTERT mRNA expression was seen in the ER-negative RL 95-2 and HEC-1B cells, regardless of E2 concentration or time of exposure. Reconstitution of ERα expression by transfection in the RL 95-2 cells led to induction of telomerase expression by E2. This demonstrates the importance of an intact ER in estrogen's ability to modulate telomerase activity in the endometrial cancer cell lines. As demonstrated by others [9,10], our findings support a direct interaction of activated ERα with two putative EREs in the

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Fig. 5. Estrogen activates hTERT promoter. (A) Schematic diagram of hTERT promoter luciferase plasmids showing two ERE binding sites and core promoter. (B) ER-positive Ishikawa cells were transfected with hTERT promoter luciferase plasmids, and luciferase activity was assayed after exposure to E2 (1 μM) for 48 h. (C) ERα-transfected RL 95-2 cells were subsequently transfected with hTERT promoter luciferase plasmids, and luciferase activity was assayed after exposure to E2 (1 μM) for 48 h. Data are presented as means ± standard deviation (SD) of duplicated samples from at least two independent experiments.

5′ flanking sequence of the hTERT promoter, leading to E2dependent stimulation of hTERT gene transcription. The distal ERE is located at − 2777/− 2755, and the proximal one at − 979/ − 956, which overlaps with an Sp1 binding site. Gel shift assays confirmed that both of these putative EREs in the hTERT promoter specifically bind to ERα as demonstrated in the ERpositive Ishikawa cell line and in the ERα-transfected RL 95-2 cell line. Transient expression assays using luciferase reporter plasmids containing various fragments of the hTERT promoter demonstrated that at least the proximal ERE is responsible for transcriptional activation by ligand-stimulated ERα. Similar half ERE/Sp1 sites have been previously shown to function as EREs in some estrogen responsive gene promoters [17,18]. The function of the distal ERE was not evaluated in this study; and thus, its role in the estrogen-induced transcriptional activation of hTERT remains unclear. Telomerase activity is thought to be integral in maintaining the physical function of the endometrium. Estrogen levels and telomerase activity have been previously shown to increase in parallel in the proliferative phase of the menstrual cycle [4]. This work supports a direct interaction between estrogen and telomerase regulation through mediation of hTERT gene transcription that may contribute to estrogen-induced endometrial

carcinogenesis. Our results demonstrating estrogen-induced telomerase activity after reconstitution of ERα in RL 95-2 cells suggest that ERα and not ERβ may be predominantly involved in the regulation of telomerase activity in endometrial cancer cells. This finding is consistent with previous work performed in breast cancer cell lines and normal ovarian epithelium [9,10]. However, a limitation to this work is that ERβ was not reconstituted in the RL 95-2, and its effect on telomerase activity was not directly evaluated. Although ERα is considered to be the dominant receptor in the endometrium, ERβ has been also implicated in the modulation of estrogenic effects [19], and therefore we cannot completely exclude its potential impact on telomerase activation. Endometrial cancer is the fourth most common cancer in the United States [12] and has been increasing in frequency secondary to an aging female population and to dietary and hormonal factors. In general, endometrial cancers are divided into endometrioid and non-endometrioid histologic subtypes, also referred to as type I and type II tumors, respectively. Type I tumors are classically thought to arise from a hyperplastic precursor from which estrogen stimulation drives its malignant transformation to an endometrioid adenocarcinoma. In contrast, type II tumors are associated with an atrophic endometrium and

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the absence of excess estrogen. Thus, our work was primarily focused on better understanding the underlying relationship between estrogen and type I endometrial cancers. Obesity, exogenous estrogen treatment for postmenopausal symptoms, especially when unopposed by cyclic progestins, as well as hormonal disturbances associated with other disease states such as diabetes mellitus and polycystic ovarian syndrome have all been identified as risk factors for the development of type I endometrial cancers. To date, very little is known regarding the molecular biology involved in endometrial cancer pathogenesis, especially in regard to the role of estrogen. Estrogen receptor status in endometrial cancer specimens has been shown to be inversely proportional to histologic grade, clinical stage and overall survival [20], and the absence of expression has been linked to an increased risk of recurrence [21,22]. Although telomerase activity is weak to absent in the normal endometrium after menopause [7], reactivation of telomerase expression is seen in over 80% of endometrial carcinomas [23–26]. This is true for both type I and type II endometrial cancer [27,28]. This suggests that estrogen-induced telomerase activity may be an early step in the malignant transformation of the endometrium, at least for type I tumors. The potential mechanism for telomerase activation in nonendometrioid adenocarcinomas is less clear. Further investigation focused on the estrogen-dependent regulation of telomerase may ultimately provide insight into novel therapeutic and preventative strategies for endometrial cancer and other hormone-dependent tumors. References [1] Stewart SA, Weinberg RA. Telomerase and human tumorigenesis. Semin Cancer Biol 2000;10(6):399–406. [2] Yokoyama Y, Takahashi Y, Shinohara A, Lian Z, Tamaya T. Telomerase activity in the female reproductive tract and neoplasms. Gynecol Oncol 1998;68(2):145–9. [3] Zheng PS, Iwasaka T, Yamasaki F, Ouchida M, Yokoyama M, Nakao Y, et al. Telomerase activity in gynecologic tumors. Gynecol Oncol 1997;64 (1):171–5. [4] Williams CD, Boggess JF, LaMarque LR, Meyer WR, Murray MJ, Fritz MA, et al. A prospective, randomized study of endometrial telomerase during the menstrual cycle. J Clin Endocrinol Metab 2001;86(8):3912–7. [5] Saito T, Schneider A, Martel N, Mizumoto H, Bulgay-Moerschel M, Kudo R, et al. Proliferation-associated regulation of telomerase activity in human endometrium and its potential implication in early cancer diagnosis. Biochem Biophys Res Commun 1997;231(3):610–4. [6] Kyo S, Kanaya T, Ishikawa H, Ueno H, Inoue M. Telomerase activity in gynecological tumors. Clin Cancer Res 1996;2(12):2023–8. [7] Kyo S, Takakura M, Kohama T, Inoue M. Telomerase activity in human endometrium. Cancer Res 1997;57(4):610–4. [8] Cong YS, Wright WE, Shay JW. Human telomerase and its regulation. Microbiol Mol Biol Rev 2002;66(3):407–25. [9] Misiti S, Nanni S, Fontemaggi G, Cong YS, Wen J, Hirte HW, et al. Induction of hTERT expression and telomerase activity by estrogens in human ovary epithelium cells. Mol Cell Biol 2000;20(11):3764–71.

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