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(American Journal of Pathology. 2003;162:1789-1806.)
© 2003 American Society for Investigative Pathology

Hypoxia-Inducible Erythropoietin Signaling in Squamous Dysplasia and Squamous Cell Carcinoma of the Uterine Cervix and Its Potential Role in Cervical Carcinogenesis and Tumor Progression

Geza Acs^*, Paul J. Zhang^*, Cindy M. McGrath^*, Peter Acs, John McBroom, Ahmed Mohyeldin, Suzhen Liu, Huasheng Lu and Ajay Verma

From the Department of Pathology and Laboratory Medicine,^* University of Pennsylvania Medical Center, Philadelphia, Pennsylvania; the Department of Medicine, Huron Hospital, Cleveland Clinic Health System, Cleveland, Ohio; the Department of Obstetrics and Gynecology, Walter Reed Army Medical Center, Washington, D.C.; and the Department of Neurology, Uniformed Services University of the Health Sciences, Bethesda, Maryland

	Abstract

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Tissue hypoxia is a characteristic property of cervical cancers that makes tumors resistant to chemo- and radiation therapy. Erythropoietin (Epo) is a hypoxia-inducible stimulator of erythropoiesis. Acting via its receptor (EpoR), Epo up-regulates bcl-2 and inhibits apoptosis of erythroid cells and rescues neurons from hypoxic damage. In addition to human papillomavirus infection, increased bcl-2 expression and decreased apoptosis are thought to play a role in the progression of cervical neoplasia. Using reverse transcriptase-polymerase chain reaction and Western blotting we showed that HeLa and SiHa cervical carcinoma cells and human cervical carcinomas express EpoR, and that hypoxia enhances EpoR expression. Exogenous Epo stimulated tyrosine phosphorylation and inhibited the cytotoxic effect of cisplatin in HeLa cervical carcinoma cells. Using immunohistochemistry, we examined the expression of Epo, EpoR, p16, hypoxia-inducible factor (HIF)-1, and bcl-2 in benign and dysplastic cervical squamous epithelia and invasive squamous cell carcinomas (ISCCs). EpoR expression in benign epithelia was confined to the basal cell layers, whereas in dysplasias it increasingly appeared in more superficial cell layers and showed a significant correlation with severity of dysplasia. Diffuse EpoR expression was found in all ISCCs. Expression of Epo and HIF-1 was increased in dysplasias compared to benign epithelia. Focal Epo and HIF-1 expression was seen near necrotic areas in ISCCs, and showed correlation in their spatial distribution. Significant correlation was found between expression of EpoR, and p16 and bcl-2 in benign and dysplastic squamous epithelia. Our results suggest that increased expression of Epo and EpoR may play a significant role in cervical carcinogenesis and tumor progression. Hypoxia-inducible Epo signaling may play a significant role in the aggressive behavior and treatment resistance of hypoxic cervical cancers.

Squamous cell carcinoma accounts for 75 to 85% of cervical cancers⁶ and develops in the background of increasing grades of dysplasia (termed cervical intraepithelial neoplasia, CIN).^6-9 High-risk human papillomavirus (HPV) DNA is detected in more than 90% of malignant cervical tumors and it is widely accepted that HPV plays an essential role in the pathogenesis of cervical cancer.^7,8,10 The high-risk HPV oncoproteins, E6 and E7, have been shown to be necessary for immortalization and transformation of cervical keratinocytes.¹¹ E6 binds to the wild-type p53 protein and promotes its ubiquitin-dependent degradation.¹² E7 binds to the retinoblastoma protein Rb, and disrupts the complex between Rb and the E2F transcription factor family, which controls the expression of genes involved in cell-cycle progression.¹³ In normal cells the activity of cyclin-dependent kinase (CDK)4 and CDK6 is tightly regulated by several cyclin-dependent kinase inhibitors including p16^INK4a.^14,15 Because expression of p16^INK4a underlies a negative feedback control through Rb,¹⁶ the reduction or loss of Rb function (through HPV E7) results in enhanced p16^INK4a levels in respective cells.¹⁷ Indeed, overexpression of p16^INK4a has been demonstrated in cervical squamous cell carcinoma and squamous dysplasia infected with high-risk HPV types.¹⁸

Despite these associations, only a small percentage of precursor lesions infected with HPV develop into invasive carcinomas.⁷ Therefore, additional genetic and microenvironmental factors subsequent to HPV infection must play an important role in the initiation and progression of cervical neoplasia.^7,8 The potential of tumor progression is thought to be associated with the balance between cell death and cell proliferation and apoptosis plays an important role in the maintenance of this balance.^19,20 Several studies support an important role for apoptosis and altered expression of the anti-apoptotic protein bcl-2 in the development and progression of human cervical carcinoma.^21-25

Hypoxic microregions are a characteristic pathophysiological property of solid tumors^26-29 and are thought to result from inadequate perfusion because of severe structural and functional abnormalities of the tumor microcirculation and from tumor-related anemia.^27,30 Previous studies have shown that 60% of locally advanced cervical squamous cell carcinomas have hypoxic tissue areas, which are heterogeneously distributed within the tumor mass.^27-29,31,32 Tumor hypoxia makes solid tumors resistant to radiation and chemotherapy.^27,33,34 More recent studies suggest that sustained hypoxia can additionally enhance local and systemic malignant progression and may increase aggressiveness through clonal selection and genomic and proteomic changes.^{27,31,32,35,36} Multivariate analysis has shown that tumor hypoxia is indeed the most powerful prognostic factor in cervix cancers independent of other variables.^33,34,37,38

Many hypoxia-regulated genes, such as vascular endothelial growth factor, are known to play a key role in carcinogenesis and tumor progression.^39,40 The best known hypoxia-regulated gene is erythropoietin (Epo), a glycoprotein hormone stimulator of erythropoiesis.^41-43 Epo gene expression is primarily modulated by tissue hypoxia^44,45 and this regulation occurs mainly at the mRNA level mediated by hypoxia-inducible transcription factor-1 (HIF-1).^41,44,46 During adult life, Epo is normally produced by the kidney and liver.^41,43,47 The Epo receptor (EpoR) belongs to the cytokine receptor type I superfamily.^41,48-54 The signaling mechanisms after receptor activation include the Jak/STAT and the Ras/MAP kinase pathways.^41,46,48-59 EpoR stimulation in erythroblasts promotes their proliferation and differentiation, and leads to increased expression of the anti-apoptotic proteins bcl-2 and bcl-X_L,^57,58 and inhibition of apoptosis.^{46,52,53,56,59}

Recently other sites of Epo production have been reported, including bone marrow macrophages,⁶⁰ brain astrocytes,^61,62 trophoblast cells of human placenta,⁶³ and the breast.^64-66 Epo production was also demonstrated in human female reproductive organs, including the uterus.^67,68 In recent years it has also become clear that EpoR is expressed by a variety of cell types, including endothelial cells,⁶⁹ neurons,⁷⁰ trophoblast cells,⁷¹ and mammary epithelial cells.^64-66 Although the specific function(s) of EpoR in these nonhematopoietic sites is not fully understood, the EpoR expressed in these tissues seems to be functional, thus suggesting a wider biological role for Epo signaling unrelated to erythropoiesis.^72,73 There appears to be a paracrine Epo/EpoR system in the brain, where neurons express EpoR^61,70,74 and astrocytes produce Epo.^61,62 Evidence suggests that the signaling cascades that have been characterized in hematopoietic cells,^48-50,52,56 are also functional in neurons and can be modulated by Epo. It has been demonstrated in vitro and in vivo that Epo is a potent inhibitor of neuronal apoptosis induced by ischemia and hypoxia.⁷⁵ EpoR mRNA is also expressed in endothelial cells and Epo stimulates proliferation and migration of human endothelial cells^76,77 and angiogenesis.⁶⁸

Elevated Epo levels have long been recognized in patients with renal cell carcinomas, Wilms’ tumors, hepatomas, and cerebellar hemangioblastomas, all tumors arising in anatomical sites in which Epo is normally expressed in low levels.⁴⁴ Moreover, ectopic Epo expression in erythroleukemia cells was found to mediate their autonomous growth.⁷⁸ EpoR expression was also reported in cases of renal cell carcinoma and a potential paracrine or autocrine role for Epo signaling for promoting growth of renal carcinomas has been suggested.⁷⁹ We have recently described that human breast cancer cell lines and human breast carcinomas express Epo and EpoR mRNA and protein and that their expression is enhanced by hypoxia.^64,65 Furthermore, we demonstrated that Epo signaling is biologically active and stimulates tyrosine phosphorylation, DNA synthesis, and proliferation in breast cancer cells.

In the present study we report biologically active, hypoxia-inducible Epo signaling in human cervical carcinoma cells. We demonstrate that human recombinant Epo (rHu-Epo) inhibits the cytotoxic effect of the chemotherapeutic drug cisplatin in cervical carcinoma cells. Using immunohistochemistry we demonstrate the expression of Epo and EpoR in human tissue samples and cytological smears of cervical squamous dysplasia and carcinoma, and correlate their expression with that of p16^INK4a, HIF-1, and bcl-2.

	Materials and Methods

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Cell Culture and Treatments

All culture media were purchased from Life Technologies, Inc. (Rockville, MD). The HeLa and SiHa human cervical carcinoma, Hep3B human hepatoma, and U251 human glioblastoma cell lines were obtained from the American Type Culture Collection (ATCC, Manassas, VA) and cultured according to ATCC directions. Forty-eight hours before hypoxic treatment cells were switched to serum-free medium. Hypoxic treatment of cells was performed in an enclosed chamber (Billups-Rothenberg Inc., Del Mar, CA) flushed with a premixed gas mixture (1% O₂, 5% CO₂, 94% N₂) for the times indicated. The O₂ concentration in the culture medium was confirmed to be 1% using an oxygen-sensitive electrode (World Precision Instruments, Sarasota, FL), and this oxygen concentration was maintained throughout the course of the experiments. Stimulation of tyrosine phosphorylation was performed by treating cells in phenol-free Dulbecco’s modified Eagle’s medium with recombinant human erythropoietin (rHu-Epo, epoetin-, 10 U/ml; Amgen, Thousand Oaks, CA) or the Epo mimetic peptide Emp1 (AEHCSLNENITVPDTKV, 1 µg/ml) for 30 minutes.

Cytotoxicity Assay

For cytotoxicity studies cells were plated in 96-well plates at 3000 cells/well. The following morning cells were treated with rHuEpo (0, 25, 50, or 200 U/ml) with or without the general tyrosine kinase inhibitor genistein (10 µmol/L) for 60 minutes. Cells were then exposed to cisplatin (10 µg/ml) for 20 hours. Each condition was performed in quadruplicate. Viability of cells was assessed using the non-radioactive cell proliferation (MTT) assay (Promega, Madison, WI).

Annexin V Apoptosis Assay

Apoptosis was assayed by detection of membrane externalization of phosphatidylserine using the Annexin V-PE Apoptosis Detection Kit I (BD Biosciences, Palo Alto, CA). Cells were plated at 40 to 50% confluency in six-well plates and allowed to adhere overnight. Media was then removed and replaced with serum-free media with or without rHuEpo. Cells were pretreated with rHuEpo (0, 50, 100, or 200 U/ml) for 2 hours and then challenged with cisplatin (5 µg/ml) for 48 hours. Both adherent and floating cells were harvested for the apoptosis assay. Cells were washed once with phosphate-buffered saline, resuspended in binding buffer, and stained with phycoerythrin-conjugated Annexin V and 7-amino-actinomycin (7-AAD) according to the manufacturer’s protocol (5 µl for 1 x 10⁵ cells) for 15 minutes. Cells were analyzed using an EPICS XL-MCL (Beckman Coulter, Miami, FL) flow cytometer. Emission from Annexin V was collected through a 575-nm band-pass filter and emission from 7-AAD was collected through a 675-nm band-pass filter. A minimum of 10,000 events was collected for each sample and each treatment group was done in triplicates.

Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR)

Measurement of Epo and EpoR mRNA were performed as described previously.^64,80 RNA was isolated using the RNeasy RNA isolation kit (Qiagen, Inc., Valencia, CA) and the Superscript One-Step RT-PCR System (Life Technologies, Inc., Gaithersburg, MD) was used for RT-PCR. Approximately 1 µg of RNA and 10 pmol of both the forward and reverse primers were used for each sample. RT was performed at 50°C for 30 minutes. The PCR cycle protocol for the Epo primers was 42 cycles at 94°C for 20 seconds, 61°C for 45 seconds, and 72°C for 1 minute, and for EpoR was 42 cycles at 94°C for 15 seconds, 55°C for 30 seconds, 72°C for 1 minute. Primers for Epo were: sense 5'-ACTCTGCTTCGGGCTCTGGGAGCCCAGAAG-3' (corresponding to nucleic acids 581 to 610), and anti-sense 5'-AAGCAATGTTGGTGAGGGAGGTGGTGGAT-3' (corresponding to nucleic acids 786 to 814).^64,80 Primers for EpoR were: sense 5'-GGCAGTGTGGACATAGTGGC-3' (corresponding to nucleic acids 1304 to 1323), and anti-sense 5'-AGCAGGATGGATTGGGCAGA-3' (corresponding to nucleic acids 1782 to 1801). These primer sets were used previously, with subsequent nucleotide sequencing of the 447-bp EpoR RT-PCR products obtained from MCF-7 human breast cancer and MG-U87 human glioblastoma cells demonstrating that these were identical to the normal human EpoR (BLAST NCBI search results).⁶⁴ The RT-PCR products were then run on a 1% agarose gel. The DNA standard used was the 100-bp DNA ladder from FMC Bioproducts (BioWhittaker Molecular Applications, Rockland, ME).

Western Blotting

For detection of EpoR, whole cell lysates and clinical biopsy sample protein extracts were normalized for protein. One hundred µg of proteins from each sample were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis, transferred to nitrocellulose, and stained with anti-EpoR antibody (rabbit polyclonal, C20, 1:1500 dilution; Santa Cruz Biotechnology, Inc., Santa Cruz, CA) in the absence or presence of EpoR blocking peptide (10:1 peptide:antibody ratio, Santa Cruz Biotechnology). Secondary antibody was goat anti-rabbit antibody conjugated with horseradish peroxidase (Amersham Pharmacia Biotech, Piscataway, NJ). Immunoreactive bands were visualized using chemiluminescence (SuperSignal WestPico Chemiluminescence kit; Pierce, Rockford, IL). A horseradish peroxidase-conjugated monoclonal antibody (PY20, SC-508, 1:1000 dilution; Santa Cruz Biotechnology) was used to detect phosphotyrosine. Nuclear extracts and Western blot analysis for HIF-1 were performed as previously described⁸¹ using a mouse monoclonal anti-HIF-1 antibody (clone 54, 1:500 dilution; BD Transduction Laboratories, San Jose, CA). Secondary antibody was rabbit anti-mouse conjugated with horseradish peroxidase (Amersham Pharmacia Biotech).

Clinical Samples and Clinical Data

Study protocols involving human material were approved by the University of Pennsylvania Institutional Review Board. Fresh tissue from seven cervical squamous cell carcinomas and adjacent benign cervical squamous mucosa (in one case) was obtained at the time of surgery. A frozen section was cut from each sample and examined to confirm the presence or absence of tumor tissue: all tumor samples contained at least 80% neoplastic cells, whereas no tumor cells were present in the sample of benign mucosa. Tissue samples were stored at -80°C until used for protein extraction.

Ninety-seven cases of cervical biopsies, cone biopsies, or hysterectomies performed for benign disease (leiomyomas), cervical dysplasia, or invasive squamous cell carcinoma (ISCC) were selected from the surgical pathology files of the University of Pennsylvania Medical Center. Hematoxylin and eosin (H&E)-stained slides of all cases were reviewed and the diagnoses confirmed. Grading of squamous dysplasias (CIN I to CIN III) was performed on the H&E-stained slides according to established criteria.⁹ Invasive carcinomas were also evaluated to determine the presence or absence of tumor cell keratinization and necrosis. Information regarding clinicopathological tumor features were retrieved from the pathology reports and are summarized in Table 1 .

Immunohistochemistry and Immunocytochemistry

Immunohistochemical assays were performed on formalin-fixed paraffin-embedded sections as described previously.⁶⁵ Five-µm-thick sections were cut and deparaffinized in xylene and rehydrated in graded alcohols. SurePath cytological slides were decoverslipped in xylene, rehydrated in graded alcohols, and postfixed in 10% phosphate-buffered formaldehyde for 30 minutes. All slides were steamed in 0.01 mol/L of sodium citrate buffer (pH 6.0) for 20 minutes. Endogenous peroxidase activity was blocked by 3% hydrogen peroxide in methanol for 20 minutes. Slides were incubated with the antibodies against Epo (rabbit polyclonal, H-162, 1:200 dilution; Santa Cruz Biotechnologies), EpoR (rabbit polyclonal, C-20, 1:400 dilution; Santa Cruz Biotechnologies), bcl-2 (clone 124, mouse monoclonal, 1:50 dilution; DAKO Corp., Carpinteria, CA), p16^INK4a (clone 16P07, mouse monoclonal, 1:100 dilution; Neomarkers, Fremont, CA), and HIF-1 (clone H1alpha67, mouse monoclonal, 1:10,000; Neomarkers) overnight at 4°C. In the cases of Epo, EpoR, bcl-2, and p16^INK4a assays, the slides were then washed five times with Tris-buffered saline containing Tween 20 (TBST, pH 7.6; DAKO) and incubated for 30 minutes at room temperature with horseradish peroxidase-labeled dextran polymer coupled to anti-rabbit or anti-mouse antibody (DAKO EnVision + System HRP, DAKO). Slides were then washed three times with TBST, developed with diaminobenzidine for 10 minutes, and counterstained with hematoxylin. For HIF-1 the DAKO CSA signal amplification kit was used according to the manufacturer’s recommendations. For Epo and EpoR, slides of fetal liver⁸² and placenta^63,71 were used as positive controls. For bcl-2 slides of human tonsil, and for p16^INK4a slides of human cervical squamous cell carcinoma known to show strong p16^INK4a immunostaining were used as positive controls. For HIF-1 slides of oral squamous cell carcinoma with large areas of necrosis known to show HIF-1 overexpression were used as positive control. A negative control was done in each case by omission of the primary antibody. The specificity of the Epo and EpoR antibodies were confirmed previously.^{64,66,70,75,83} In addition, the specificity of the EpoR and Epo immunoreactivity was also evaluated by antibody absorption test: the primary antibody was preincubated with blocking peptide for EpoR (Santa Cruz Biotechnologies Inc.) or human recombinant Epo (rHuEpo; R&D Systems, Minneapolis, MN) (10:1 peptide:antibody ratio), which resulted in complete abolishment of immunohistochemical staining. The specificity of the immunostaining reaction is further supported by other experiments using a mouse monoclonal anti-Epo (clone 9C21D11, R&D Systems) and a rabbit polyclonal anti-EpoR antibody (Upstate Biotechnology, Lake Placid, NY),⁸⁴ which resulted in an immunostaining pattern similar to that obtained with the antibodies used in the current study (Acs et al, unpublished observation).

Double Immunohistochemistry for HIF-1 and Epo

For double-immunohistochemical staining slides were immunostained for HIF-1 using the DAKO CSA kit as described above. After incubation with diaminobenzidine, the slides were washed three times in deionized water and three times in TBST. Slides were then incubated with the primary Epo antibody as described above, washed five times with TBST, and incubated for 30 minutes at room temperature with alkaline phosphatase-labeled dextran polymer coupled to anti-rabbit antibody (DAKO EnVision + System AP, DAKO). Slides were then washed three times with TBST, developed with fast red chromogen for 10 minutes, and counterstained with hematoxylin.

Interpretation of Immunohistochemical Stains

Immunohistochemical stains for Epo, EpoR, p16^INK4a, and HIF-1 were interpreted semiquantitatively by assessing the intensity and extent of staining on the entire tissue sections present on the slides according to a four-tiered (0 to 3) scale. For Epo cytoplasmic, and for EpoR cytoplasmic and/or membrane immunoreactivity was considered positive. For HIF-1 nuclear staining, and for p16^INK4a nuclear and/or cytoplasmic staining was considered positive. In the case of dysplasias or in situ carcinomas, first, the percentage of total epithelial thickness showing positive staining was determined (eg, 50% if the basal half or 75% if the basal three-fourths of the squamous epithelium showed positive immunostaining, and so forth). In the case of invasive tumors, first the total percentage of positively staining tumor cells was determined. Then the percentage of weakly (1), moderately (2), and strongly (3) staining cells was determined, so that the sum of these categories equated with the overall percentage of positivity. A staining score was then calculated as follows: score (out of maximum of 300) = sum of 1 x percentage of weak, 2 x percentage of moderate, and 3 x percentage of strong staining. To assess the immunostaining for Epo and HIF-1 adjacent to areas of tumor cell necrosis and keratinization, immunostaining was scored as described above in at least 10 high-power fields of viable tumor cells within 1 mm of necrotic or keratinized areas, and in tumor regions away from such areas. Bcl-2-stained slides were interpreted by determining the percentage of squamous epithelial cells showing cytoplasmic immunoreactivity of any intensity. Immunohistochemical stains were evaluated independently by two pathologists (GA and PJZ). Slight differences in interpretation were resolved by simultaneous viewing.

Immunofluorescence

For immunofluorescence staining slides were pretreated and incubated with the primary antibody against p16^INK4a or EpoR as described above. Slides were then washed five times with TBST and incubated with a fluorescein isothiocyanate-conjugated rabbit anti-mouse secondary antibody (1:20 dilution, DAKO) or a tetramethyl-rhodamine isothiocyanate-conjugated swine anti-rabbit secondary antibody (1:40 dilution, DAKO) for 30 minutes on 37°C in the case of p16^INK4a and EpoR, respectively. Slides were then coverslipped with 4,6-diamidino-2-phenylindole and examined using a Leica DMRA Research Microscope equipped with a Megapixel charge-coupled device camera. For simultaneous immunostaining of tissue sections and cytological preparations for p16^INK4a and EpoR, slides were first stained for EpoR as described above. Slides were then washed five times with TBST and immunostained for p16^INK4a as described above.

Statistical Analysis

The Wilcoxon signed rank test was used for the comparison of median p16^INK4a, EpoR, Epo, HIF-1, and bcl-2 immunohistochemical expression levels in dysplastic epithelia and the adjacent benign squamous epithelium, and for the comparison of Epo and HIF-1 immunostaining in tumor areas adjacent to or away from necrosis and keratinization. Median p16^INK4a, EpoR, Epo, HIF-1, and bcl-2 immunohistochemical expression levels in benign epithelia, CIN I, CIN II, CIN III, and ISCC were compared using the Kruskal-Wallis one-way analysis of variance by ranks followed by Dunn’s multiple comparison test, when appropriate. The correlation between the levels of Epo and EpoR staining and p16, bcl-2, and HIF-1 staining was estimated using the Spearman rank correlation test. The normal versus abnormal distribution of HIF-1 expression in benign and dysplastic epithelia was compared using the Fisher’s exact test. Statistical significance was determined if the two-sided P value of a test was less than 0.05.

	Results

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Hypoxia-Inducible Expression of Functional EpoR in Human Cervical Carcinoma Cell Lines and Human Cervical Carcinomas

Using RT-PCR, HeLa and SiHa human cervical carcinoma cells cultured under normoxia (21% O₂) or hypoxia (1% O₂) were analyzed for Epo and EpoR mRNA expression. Abundant expression of EpoR mRNA was detected in HeLa cells under both conditions, whereas SiHa cells showed a prominent increase in EpoR mRNA after hypoxia treatment (Figure 1A) . No Epo mRNA expression could be detected in HeLa and SiHa cervical carcinoma cells (Figure 1A) . In contrast, human hepatoma Hep3B and glioblastoma U251 cells expressed abundant Epo mRNA (Figure 1B) , as reported previously.⁶⁴

View larger version (64K): [in this window] [in a new window]   Figure 1. RT-PCR analysis of Epo and EpoR expression in HeLa and SiHa cervical carcinoma cells. A: HeLa and SiHa cells cultured under normoxia (N, 21% O2) or hypoxia (H, 1% O2) were analyzed for Epo and EpoR mRNA expression. Although abundant expression of EpoR mRNA was detected in HeLa cells under both conditions, SiHa cervical carcinoma cells showed a prominent increase in EPOR mRNA expression after hypoxia treatment. Epo mRNA expression could not be detected in either cell lines. B: In contrast, human hepatoma Hep3B and glioblastoma U251 cells expressed abundant Epo mRNA.

View larger version (54K): [in this window] [in a new window]   Figure 2. HeLa cervical carcinoma cells express hypoxia-inducible and functional EpoR. A: Hypoxia activates HIF-1 and stimulates EpoR expression in cervical carcinoma cells. HeLa cells were cultured under normoxia (N, 21% O2) or hypoxia (H, 1% O2) for 3 hours and then their nuclear extracts examined by Western blotting for HIF-1 (HIF) immunoreactivity. For evaluation of EpoR expression, membranes from HeLa cells cultured under normoxia (N) or hypoxia (H) for 24 hours were used. HeLa cells cultured under normoxia (N) express a strongly immunoreactive band at Mr 66,000 identified with the antibody. Under normoxic culture conditions HeLa cells show very low levels of HIF-1 expression. Hypoxia (H) for 3 hours dramatically increased the level of HIF-1 protein expression and resulted in an increase in EpoR protein expression after 24 hours. B: EpoR-mediated activation of tyrosine phosphorylation in HeLa cells. HeLa cells were treated with either 10 U/ml of rHuEpo (Epo) or 1 µg/ml of Epo-mimetic peptide (Emp1) for 30 minutes and then analyzed for tyrosine phosphorylation by Western blotting. Phosphotyrosine immunoreactivity was dramatically enhanced by both Epo and Emp1 with the most prominent effect being at a molecular weight of 120 to 130 kd. C: Western blot analysis of cervical tissue samples (1, benign squamous mucosa; 2 to 8, cervical carcinoma; 9, HeLa cervical carcinoma cells) in the absence (-pep) or presence (+ pep) of EpoR-blocking peptide. In the absence of the blocking peptide a prominent immunoreactive band at Mr 66,000 corresponds to the EpoR protein in the tumor samples. No EpoR expression is seen in the sample of benign mucosa.1 Note the absence of the immunoreactive band at Mr 66,000 in the presence of the blocking peptide.

The majority of chemotherapy regimens used to treat patients with cervical carcinoma includes cisplatin.^85-87 Given the potent anti-apoptotic effect of Epo^{46,52,53,56,59} , we examined whether EpoR signaling may inhibit cisplatin-induced cytotoxicity and apoptosis in cervical carcinoma cells. We treated HeLa cells with cisplatin in the absence and presence of rHuEpo. Pretreatment of HeLa cells with rHuEpo showed a dose-dependent protective effect on HeLa cells treated with the chemotherapeutic agent cisplatin, as determined by the MTT assay (Figure 3A) . Using the annexin V assay, 10% of the cell killing caused by cisplatin was purely apoptotic, and rHuEpo dose dependently inhibited this apoptosis (Figure 3B) .

View larger version (19K): [in this window] [in a new window]   Figure 3. rHuEpo reverses toxicity of the chemotherapeutic drug cis-platin in cervical carcinoma cells. A: HeLa cells were treated with rHuEpo in the presence or absence of the general tyrosine kinase inhibitor genistein (10 µmol/L) for 1 hour. Cells were then exposed to cis-platin (10 µg/ml) for 20 hours. Cell viability was assessed using the MTT assay. Results show the fraction of surviving cells versus cells not treated with cis-platin. Epoetin dose dependently inhibited cis-platin cytotoxicity and this effect was significantly diminished by genistein. B: Cis-platin-induced apoptosis in HeLa cells was examined by detection of membrane externalization of phosphatidylserine using the Annexin V assay. HeLa cells were pretreated with rHuEpo for 2 hours and then challenged with cis-platin (5 µg/ml) for 48 hours. Results are expressed as percentage of the apoptotic cell population compared to the control. Cis-platin-induced apoptosis was dose dependently reversed by rHuEpo (*, P < 0.05).

To further investigate the potential role of Epo signaling in cervical squamous dysplasia and carcinoma we examined the expression of Epo and EpoR by immunohistochemistry in a series of clinical samples of cervical dysplasia and carcinoma and compared their expression with those of p16^INK4a, HIF-1, and bcl-2. The results of the immunohistochemical assays are summarized in Table 2 and illustrated in Figures 4 and 5 .

View this table: [in this window] [in a new window]   Table 2. Expression of p16, EpoR, Epo, HIF1-, and bcl-2 in Benign Squamous Epithelium, Squamous Dysplasia, and Invasive Squamous Cell Carcinoma

View larger version (20K): [in this window] [in a new window]   Figure 4. Expression of p16INK4a (p16) (A), EpoR (B), Epo (C), and HIF-1 (D) in benign cervical squamous epithelium, cervical dysplasia (CIN, grades I to III), and ISCC, determined by immunohistochemistry. Bars indicate median immunostaining score values.

Cytoplasmic and/or membrane immunostaining for EpoR was present in all samples (Table 2 , Figures 4B and 5 ). In benign squamous epithelia EpoR expression was confined to the basal and parabasal cell layers. In contrast, in squamous dysplasia EpoR expression increasingly appeared in the more superficial cell layers and was significantly increased compared to the adjacent benign epithelia (P = 0.0002 for CIN I, P = 0.0003 for CIN II, and P < 0.0001 for CIN III; Wilcoxon signed rank test). In CIN I and CIN II cases EpoR expression was seen in the basal 50% and 70% of the epithelial thickness, whereas in CIN III the full thickness of the dysplastic epithelia showed EpoR expression (Figure 6) . The mean EpoR staining score increased with increasing grade of dysplasia (slope 51.72 ± 4.61, r² = 0.9844, P = 0.0078, linear regression). Prominent EpoR expression was also present in the endothelial and smooth muscle cells of blood vessels (not shown). Immunofluorescence staining for EpoR resulted in a similar immunostaining pattern, including prominent labeling of endothelial and smooth muscle cells of blood vessels (Figure 6C) .

View larger version (70K): [in this window] [in a new window]   Figure 6. Expression of EpoR increases with increasing severity of dysplasia in cervical squamous epithelia. A: Immunohistochemical expression of EpoR in benign squamous epithelium and CIN grades I, II and III. In benign squamous epithelium EpoR expression is seen only in the basal one or two cell layers. With increasing severity of dysplasia EpoR immunostaining appears in the more superficial cell layers, and in CIN III, EpoR immunostaining is present in full thickness of the epithelium (immunohistochemical stains for EpoR with hematoxylin counterstain; H&E). B: EpoR expression increases with increasing severity of dysplasia in cervical squamous epithelia. Data are mean values ± SEM of percent epithelium thickness showing EpoR expression. C: Strong EpoR expression in endothelial and smooth muscle cells of a small artery and a lymphatic channel (arrowhead) (immunofluorescence staining for EpoR, tetramethyl-rhodamine isothiocyanate, red; nuclei are stained with 4,6-diamidino-2-phenylindole, blue; original magnification, x400).

View larger version (179K): [in this window] [in a new window]   Figure 7. Expression of Epo and HIF-1 in cervical squamous cell carcinomas show correlation in spatial distribution. Prominent Epo (A) and HIF-1 (C) expression are present in tumor cells adjacent to necrotic areas (asterisk). Epo (B) and HIF-1 (D) expression is also present adjacent to tumor areas showing tumor cell keratinization. Double-immunohistochemical stains (E and F) show co-expression of Epo (cytoplasmic staining, red) and HIF-1 (nuclear staining, brown) in cervical carcinomas adjacent to necrotic areas (asterisk) (immunohistochemical stains with hematoxylin counterstain).

View this table: [in this window] [in a new window]   Table 3. Erythropoietin (Epo) and Hypoxia-Inducible Factor 1- (HIF) Immunostaining in Invasive Squamous Cell Carcinoma of the Cervix in Areas of Tumor Cell Necrosis and Keratinization

View larger version (34K): [in this window] [in a new window]   Figure 8. Distribution of HIF-1 immunostaining in benign and dysplastic cervical squamous epithelia. Basal, mid, and upper designate the basal, mid, and upper thirds of the squamous epithelium. Each bar represents one case and indicates the presence of immunostaining in the respective portion of the squamous epithelium. Note that in benign squamous epithelia HIF-1 immunostaining is primarily restricted to the suprabasal, middle portion of the epithelium. In contrast, in most cases of dysplasia HIF-1 immunostaining showed an abnormal distribution and was present in the basal and/or more superficial epithelial cells as well.

View larger version (135K): [in this window] [in a new window]   Figure 5. Immunohistochemical expression of p16INK4a (p16), EpoR, Epo, HIF-1, and bcl-2 in benign cervical squamous epithelium (benign), low-grade squamous dysplasia (CIN I), high-grade squamous dysplasia (CIN III), and ISCC (immunohistochemical stains with hematoxylin counterstain; H&E).

View larger version (17K): [in this window] [in a new window]   Figure 9. Correlation of immunohistochemical expression of EpoR with p16INK4a (p16) and bcl-2 in benign and dysplasic cervical squamous epithelia. A: The level of EpoR expression shows positive correlation with the expression of p16 (r = 0.8644, P < 0.0001, Spearman’s test). The line represents the calculated regression line. B: The level of EpoR expression shows positive correlation with the expression of bcl-2 (r = 0.4632, P < 0.0001, Spearman’s test). The line represents the calculated regression line.

View larger version (104K): [in this window] [in a new window]   Figure 10. Cells of cervical squamous dysplasia (CIN III) and ISCC show co-expression of EpoR (red) and p16INK4a (p16, green). Note that benign squamous epithelium (asterisk) shows weak EpoR expression in the basal cell layers, but no expression of p16INK4a. (Double-immunofluorescence stain for EpoR, tetramethyl-rhodamine isothiocyanate, red, and p16INK4a, fluorescein isothiocyanate, green; nuclei are stained with 4,6-diamidino-2-phenylindole, blue; H&E.)

Cytoplasmic EpoR immunostaining was seen in benign squamous cells in 2 of 39 cases. In contrast, dysplastic squamous cells showed strong cytoplasmic EpoR expression in all 16 high-grade squamous intraepithelial lesion and 4 of 5 low-grade squamous intraepithelial lesion cases (Figure 11) . The difference in EpoR expression between benign and dysplastic squamous cells was highly significant (P < 0.0001, Fisher’s exact test). Double-immunofluorescence staining showed co-expression of EpoR and p16^INK4a in dysplastic squamous cells, whereas benign squamous cells showed no staining with either antibody (Figure 11) .

View larger version (37K): [in this window] [in a new window]   Figure 11. Expression of EpoR and p16INK4a in dysplastic cervical squamous cells in SurePath liquid-based cervicovaginal cytological preparations. A, B, and E: Papanicolaou-stained preparations showing high-grade squamous intraepithelial lesion and background benign squamous cells. C and D: Cells of high-grade squamous intraepithelial lesion show prominent EpoR expression, whereas no immunostaining is seen in benign squamous cells (immunohistochemical stain for EpoR with hematoxylin counterstain). F to J: Cells of high-grade squamous intraepithelial lesion co-express p16INK4a (F and I, green), and EpoR (G and J, red). H: Triple band pass filter image showing the co-expression of both proteins in the same cells (yellow). (Double-immunofluorescence stain for EpoR, tetramethyl-rhodamine isothiocyanate, red, and p16INK4a, fluorescein isothiocyanate, green; nuclei are stained with 4,6-diamidino-2-phenylindole, blue.)

	Discussion

Top Abstract Materials and Methods Results Discussion References

We found a highly significant correlation between the expression of EpoR and overexpression of p16^INK4a in the samples. Previous studies by Klaes and colleagues¹⁷ indicate that diffuse and strong overexpression of the p16^INK4a protein is observed in almost 100% of high-grade dysplastic cervical lesions, whereas normal cervical epithelium or inflammatory and metaplastic lesions were not stained. It was suggested that p16^INK4a is a specific marker for cells that express high-risk HPV E6/E7 oncogenes and retain the capacity to replicate. In our study, the sensitivity, specificity, positive and negative predictive value of p16^INK4a overexpression for detecting squamous dysplasia (all grades combined) were 92.9%, 100%, 100%, and 93.4%, respectively. The corresponding values for the detection of high-grade squamous dysplasia (CIN II and CIN III) were 100%, 100%, 100%, and 100%, respectively. These results are in excellent agreement with those of Klaes and colleagues,¹⁷ and indicate that p16^INK4a overexpression can be used as a useful diagnostic tool in the detection of cervical dysplasia. The co-expression of EpoR and p16^INK4a in neoplastic squamous cells, also demonstrated by double-immunofluorescence staining, suggests that detection of increased EpoR expression may also serve as a useful diagnostic marker for neoplastic cervical squamous cells.

In contrast to breast^64,65 and head and neck squamous cell carcinoma cells (Cohen et al, unpublished observations), HeLa and SiHa cervical carcinoma cells did not show Epo expression by RT-PCR analysis. However, focal Epo expression was detected by immunohistochemistry in a proportion of benign and neoplastic squamous epithelia. Epo immunostaining was focal and restricted to a small minority of the cells in most cases. In benign squamous epithelia, when present, Epo staining was strictly restricted to the basal cell layer. Epo expression was significantly increased in squamous dysplasia compared to benign epithelia. Focal Epo expression was found in most ISCC cases, with the immunostaining localized to small areas surrounding necrotic regions, areas of tumor cell keratinization, and to the infiltrating edge of tumors.

Many hypoxia-inducible genes, including that of Epo, are controlled by the heterodimeric transcription factor HIF-1. Distribution of Epo immunostaining was very similar to that of the HIF-1 subunit in the cervical tumors and co-expression of Epo and HIF-1 was also demonstrated by double-immunohistochemical stains, consistent with the well-known regulation of Epo expression by HIF-1.^{39,43,44,46,88} It is thought that the principal determinant of HIF-1 accumulation in tumor cells is tissue hypoxia.⁸⁹ In human cervical carcinoma xenografts a positive correlation between HIF1- expression and oxygenation status (as determined by EF-5 binding) was found,⁹⁰ suggesting that HIF-1 expression can be used as a surrogate marker in paraffin-embedded tissue specimens for the detection of hypoxic tumor areas. Indeed, immunostaining for this protein was mainly encountered adjacent to necrotic regions at some distance from tumor vessels and at the infiltrating edge of cancers, areas thought to represent the most hypoxic parts of solid tumors.^30,91,92

However, in two cases HIF-1 immunoreactivity was observed diffusely throughout the entire tumor tissue, a finding that may be because of alternative regulatory modes of HIF-1 expression. It has been demonstrated that HIF-1 expression can in fact be induced by signals other than hypoxia. Mutations that activate oncogenes (v-src, ras) and those that inactivate tumor suppressor genes (PTEN, VHL, p53) result in increased HIF-1 expression.⁹³ Thus, although increased HIF-1 can be attributed in part to tumor hypoxia, the basal level of HIF-1 expression in tumor cells may also be markedly increased as a consequence of somatic mutations that activate oncogene products or inactivate tumor suppressors.⁹³ The increased and abnormal HIF-1 expression observed in squamous dysplasias may also be because of similar mechanisms, such as inactivation of p53 by the HPV E6 protein.^10,12 Our results and previous reports of increased HIF-1 expression in other early neoplastic lesions (such as ductal carcinoma in situ of the breast, colonic adenoma, and prostatic intraepithelial neoplasia),⁹² suggest that increased HIF-1 expression can occur early in tumor progression.

In contrast to the focal and heterogeneous staining for Epo, EpoR immunostaining was diffuse and uniform throughout the tumor tissue, with further accentuation near necrotic areas. These findings are similar to those we previously described in human breast carcinomas,⁶⁵ and suggest that although hypoxia induces EpoR expression, other oncogenic mechanisms are likely to play a role in increased EpoR expression in neoplastic cervical squamous cells.

Recently, binding of the hypoxia marker pimonidazole to suprabasal cells in normal epithelia was described and it was hypothesized that hypoxia may act as a morphogen, inducing terminal differentiation of cells.⁹⁴ Our observation of prominent HIF-1 expression in the suprabasal cells of benign squamous epithelia also supports this hypothesis. This phenomenon appears to have a counterpart in squamous cell carcinoma, in which squamous differentiation was constantly seen in tumor areas several cell layers remote from adjacent vessels.³¹ Our observation, that in squamous cell carcinomas HIF-1 expression can be consistently found adjacent to areas of squamous differentiation with keratinization, also suggest a potential role for HIF-1 in the differentiation of neoplastic squamous cells. In dysplastic squamous epithelia HIF-1 expression was not only increased, but also showed an abnormal distribution compared to benign epithelia, suggesting a potential role in the disturbed maturation of dysplastic squamous cells. At present, the molecular mechanisms of hypoxia induced terminal differentiation are primarily unknown.

Although the fundamental role of high-risk HPV infection in the pathogenesis of cervical carcinoma is well established, other (both genetic and microenvironmental) factors are thought to play a role in cervical carcinogenesis.^7,10 The anti-apoptotic protein bcl-2 is expressed in the basal cells of cervical squamous epithelia and is thought to reflect the stem cell function of this compartment. Previous studies have demonstrated increased bcl-2 expression and decreased apoptosis in cervical squamous dysplasias. With the progression of CIN, increasing levels of bcl-2 immunoreactivity were observed: the number of bcl-2-positive cells increased and immunostaining became increasingly apparent in the higher cell compartments.^23-25,95 Our current findings of increased bcl-2 expression in dysplastic squamous epithelia are in agreement with the results of these previous studies.

In erythroblasts EpoR activation results in the stimulation of proliferation, leads to increased expression of the anti-apoptotic proteins bcl-2 and bcl-X_L,^57,58 and inhibition of apoptosis.^{46,52-54,56,59,96} Because EpoR expressed by human cervical carcinoma cells seems to be functional, our results suggest that via similar mechanisms, activation of EpoR expressed in dysplastic and neoplastic cervical squamous cells may up-regulate bcl-2 expression and reduce apoptotic potential. This hypothesis is also supported by the significant positive correlation between EpoR and bcl-2 expression in cervical squamous dysplasias.

Experimental data suggest that one of the major modes of action of most anti-cancer treatment methods, including radiation and chemotherapy, may be via activation of apoptosis in sensitive cells,^19,97-104 and the tendency of a cancer cell to undergo apoptosis may have important implications for tumor progression and response to treatment.^19,99,100 Bcl-2 overexpression protects cancer cells from apoptotic cell death induced by a variety of stimuli, including radiation and most cytotoxic drugs.^104-110

In erythroblasts and neurons the major action of Epo is inhibition of apoptosis.^{46,52-54,56,59,75,96} We have shown that Epo dose dependently inhibits the cytotoxic effect and apoptosis induced by the chemotherapeutic drug cisplatin in cervical carcinoma cells, providing further evidence that Epo signaling is biologically active in cervical cancer cells. Because human cervical carcinoma cells express functional EpoR, we hypothesize that the action of Epo in cervical cancer cells is likely mediated by mechanisms similar to those described in erythroblasts and neurons. Although we have found increased bcl-2 expression in approximately half of the cervical carcinomas examined, the number of ISCC cases included in this study did not allow a meaningful analysis of a possible correlation between Epo and EpoR, and bcl-2 expression in the tumors.

A recent study by Westphal and colleagues¹¹¹ suggested that although various cancer cell lines expressed EpoR, Epo signaling was not biologically active, and Epo was not essential for the growth of the cells in culture. The findings reported in this study are in contrast with several other reports of biologically active Epo signaling in malignant cells, including cells of erythroleukemia,⁷⁸ renal cell carcinoma,⁷⁹ breast cancer,^64,112 and cancers of the female genital tract.¹¹³ The lack of biological activity of Epo in the study of Westphal and colleagues¹¹¹ may be because of methodological differences or differences in the cell lines used.

In solid tumors heterogeneously distributed areas with very low-oxygen partial pressures exist, predominantly because of a deteriorating diffusion geometry, severe structural abnormalities of tumor microvessels, disturbed microcirculation, and tumor-related anemia.^30,32 Tumor hypoxia has been traditionally considered to be a therapeutic problem because it makes solid tumors resistant to radiation and chemotherapy.²⁷ Patients with hypoxic tumors have a significantly shorter recurrence-free and overall survival, and tumor oxygenation has been shown to be the strongest independent prognostic factor in cervix cancers followed by International Federation of Gynecology and Obstetrics (FIGO) stage.^33,34,37,38 Importantly, disadvantage in outcome for hypoxic tumors is independent of the mode of primary treatment (radiation or radical surgery). Thus, it was suggested that tumor hypoxia may not just counteract oxygen-dependent therapy forms, but, through clonal selection and genomic and proteomic changes, it may also increase aggressiveness and advance tumor progression per se.^31-33

One insight into why low-oxygen conditions may affect the aggressiveness of cervical carcinomas is that in vitro hypoxia initiates the induction of apoptosis in HPV-infected, oncogenically transformed cells¹¹⁴ and acts to select for populations of cells with reduced apoptotic sensitivity.¹¹⁵ Kim and colleagues¹¹⁴ showed that in vitro, low-oxygen conditions can select for populations of HPV-infected cervical epithelial cells that have lost their sensitivity to hypoxia-induced apoptosis, a situation that may be reflected in the malignant progression of cervical carcinomas. This hypoxia-induced selection hypothesis is also consistent with the lack of apoptotic sensitivity of cervical carcinoma cell lines to low-oxygen conditions.¹¹⁴ Although at present, the mechanism of hypoxia-induced loss of apoptotic sensitivity during cell culture is unclear, evidence suggests that transformed cells that possess mutations in their apoptotic program because of inactivation of p53 or overexpression of anti-apoptotic genes have a survival advantage in a low-oxygen environment.¹¹⁴ The secondary alterations in the cellular apoptotic programs that make HPV-infected cervical epithelial cells refractory to hypoxia-induced apoptosis appear to be independent or downstream of p53, and it was suggested that genes such as bcl-2 may play a more important role in the development of their reduced apoptotic sensitivity.¹¹⁴ In fact, studies indicate that bcl-2 protein levels are increased in both CIN²⁴ and in cervical carcinoma cell lines.¹¹⁶ Our current results suggest that stimulation of EpoR signaling by hypoxia may be an important mechanism leading to increased bcl-2 expression and decreased apoptotic potential in cervical cancer cells. Further studies are needed to confirm this hypothesis.

Anemia has been a well-recognized complication of cancer and cancer treatment.¹¹⁷ One important consequence of anemia is the resulting tumor hypoxia.¹¹⁸ Although the relationship among anemia, hypoxia, transfusion, and treatment outcome is complex, anemia has traditionally been considered to be one of the most powerful prognostic factors in patients with cancer of the cervix.¹¹⁹ rHuEpo has been shown to effectively increase hemoglobin levels and is often used in patients receiving curative radio- and chemotherapy.^120,121 However, Epo is a potent growth factor that may stimulate proliferation and inhibit apoptosis of EpoR-bearing tumor cells.^64,79 Epo also stimulates proliferation and migration of vascular endothelial cells^69,77 and augments angiogenesis.^68,122-124 We have shown previously⁶⁴ and in the current study that the vasculature of solid tumors express EpoR and thus the possible detrimental effect of Epo on tumor growth may be further aggravated by its known angiogenic activity.^{69,77,122,123} Previous studies in renal cell carcinoma patients suggested that increased serum Epo levels in the absence of polycythemia carried a worse prognosis and indicated a higher incidence of progressive metastatic disease.¹²⁵ Furthermore, Epo administration to a patient with multiple myeloma may have caused further malignant transformation resulting in plasma cell leukemia called into question the safety of Epo treatment for patients with EpoR-expressing myeloma cells.¹²⁶

Although previous studies have indicated that patients treated by rHuEpo respond better to treatment and have better prognosis compared to untreated control patients with significantly lower hemoglobin levels,¹¹⁷ these studies only suggested that raising hemoglobin concentration and decreasing anemia probably have some beneficial effect in cancer patients. In addition, most of the studies have not been well controlled, and their results have recently been challenged.¹¹⁹ We are not aware of any study in the literature that compared response to curative radio/chemotherapy and outcome in cancer patients treated with rHuEpo with a control group of patients with tumors of similar type, size, grade, and treatment, whose hemoglobin levels were maintained at comparable levels by other means, such as transfusion. Currently, an international, multicenter, randomized trial is being developed to evaluate the effect of raising hemoglobin with rHuEpo versus blood transfusions in anemic patients with advanced cervical carcinoma who are receiving concurrent platinum/radiotherapy.¹²⁷

The cellular responses to Epo may collectively promote growth of EpoR-bearing tumors, and these actions may be further enhanced by either high endogenous Epo production or by exogenous Epo administration. Thus, until it is demonstrated that pharmacological doses of Epo lacks such trophic effects in vivo, we suggest that treatment of cancer patients with rHuEpo should probably be performed with some degree of caution.

In summary, we have shown that dysplastic and neoplastic human cervical squamous cells show increased expression of functional EpoR, which can be further enhanced by hypoxia. Increased expression of EpoR may lead to elevated levels of bcl-2 and decreased apoptosis in the tumor cells. Administration of Epo inhibited cisplatin-induced cytotoxicity and apoptosis in cervical cancer cells. Our results suggest that increased EpoR signaling may play an important role in cervical carcinogenesis, tumor progression, and the aggressive behavior and therapy resistance of hypoxic cervical cancers.

	Footnotes

 
Address reprint requests to Geza Acs, M.D., Ph.D., Department of Pathology and Laboratory Medicine, University of Pennsylvania Medical Center, 6 Founders Pavilion, 3400 Spruce St., Philadelphia, PA 19104. E-mail: geza{at}mail.med.upenn.edu

Supported by the McCabe Award (to G. A.), the National Institutes of Health (grant NS37814 to A. V.), and the Mary Kay Ash Charitable Foundation (to A. V.).

Accepted for publication February 13, 2003.

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