Erythropoietin is a JAK2 and ERK1/2 effector that can promote renal tumor cell proliferation under hypoxic conditions

Commercial tissue microarrays (TMA) (MC5003a, US Biomax, Inc., Rockville, MD) constructed from clinical samples obtained from a cohort of 500 patients (400 malignant tissues and 100 benign tissues from 20 different organs) were examined by immunohistochemical staining. The clinicopathologic variables of the study cohort are available at http://​www.​biomax.​us/​tissue-arrays/​Multiple_​Organ/​MC5003a. TMAs were examined by H&E for histological verification of disease status. TMAs were deparaffinized followed by antigen retrieval using citric acid buffer (pH 6.0, 95°C for 20 mins). Slides were treated with 1% hydrogen peroxide in methanol to block endogenous peroxidase activity. After 20 mins of blocking in 1% bovine serum albumin (BSA), the TMAs were incubated overnight at 4°C with anti-human EPO antibody (sc-7956; rabbit polyclonal, dilution 1/200 in 1% BSA) and anti-human EPOR antibody (sc-695; rabbit polyclonal, dilution 1/100 in 1% BSA) from Santa Cruz Biotechnology (Santa Cruz, CA). Next, the slides were incubated with 2 μg/mL of biotinylated anti-rabbit IgG secondary antibody (Vector Laboratories, Burlingame, CA) for 30 mins at room temperature. Subsequently, the sections were stained using Standard Ultra-Sensitive ABC Peroxidase Staining kit (Pierce/Thermo Fisher Scientific, San Jose, CA) and 3, 3'- diaminobenzidine (DAB; Vector Laboratories), counterstained by hematoxyline, dehydrated, and mounted with a cover slide. Mouse xenograft tumors from the human renal cancer cell line Caki-1, known to stain strongly for EPO and EPOR were used as a positive control.

The proportion of positive cells was scored by two investigators (AL, MM) in four grades and represented the estimated proportion of immunoreactive cells (0 = 0% of cells; 1 = 1% to 40%; 2 = 41% to 75% and 3 = 76% to 100%). The intensity was scored and represented the average intensity of immunopositive cells (0 = none; 1 = weak; 2 = intermediate and 3 = strong). The proportion and intensity scores were combined to obtain a total EPO or EPOR staining score, which ranged from 0 to 6. The EPO or EPOR expression level was determined based on the total EPO or EPOR staining score as follows: none = 0, low = 1 or 2, moderate = 3 or 4, high = 5 or 6 [15]. A third investigator (CJR) reviewed discrepancies and rendered a final score. The comparison between EPO and EPOR expression in human tumors and benign tissues was calculated using Mann–Whitney U test.

Human renal cancer cell lines; Caki-1, 786-O, 769-P (ATCC, Manassas, VA), and the normal primary human renal tubule epithelial cells (RPTEC; Lonza, Walkersville, MD) were available for analysis. Cancer cell lines were maintained in RPMI1640 medium supplemented with 10% fetal bovine serum, 50 units/ml penicillin and 50 mg/ml streptomycin (Invitrogen Corporation, Carlsbad, CA). RPTEC was maintained in renal epithelial cell basal medium (REBM) supplemented with REGM complex (Lonza CC-3190). All cells were incubated in humidified atmosphere at 37°C in air with 5% CO₂ (normoxic conditions). For hypoxic conditions, cells were incubated at 37°C containing 1% O₂, 5% CO₂, and balance N₂ in a humidified incubator. The oxygen level was automatically maintained with an oxygen controller (ProOx P110; Biospherix, Redfield, NY) supplied with compressed nitrogen gas. Recombinant human EPO (rhEPO) was purchased from R&D Systems, Inc. (Minneapolis, MN).

Whole cell lysates were prepared using RIPA buffer with Halt Protease Inhibitor Cocktail (Thermo Fisher Scientific) as previously reported [16]. Twenty micrograms of total protein (assessed using BCA protein assay) were subjected to SDS-PAGE using Mini-PROTEAN TGX precast gels (Bio-Rad Laboratories, Richmond, CA). Proteins were transferred to polyvinylidene difluoride (PVDF) membrane (Bio-Rad). Anti-human pVHL (#2738, dilution 1:1 000), HIF-2α (#7096, dilution 1:1 000), p-Jak2 (#4406, dilution 1:1 000), total Jak2 (#3230, dilution 1:1 000), p-Stat5 (#9359, dilution 1:1 000), total Stat5 (#9363, dilution 1:1 000), p-Akt (#4060, dilution 1:1 000), total Akt (#9272, dilution 1:1 000), p-ERK1/2 (#4370, dilution 1:1 000), total ERK1/2 (#9102, dilution 1:1 000), cyclin D1 (#2978, dilution 1:1000), cyclin D3 (#2936, dilution 1:1 000), CDK4 (#2906, dilution 1:1 000), CDK6 (#3136, dilution 1:1 000), p21^cip1 (#2947, dilution 1:1000), p27^kip1 (#3686, dilution 1:1 000) and p15 (#4822, dilution 1:1 000) were purchased from Cell Signaling Technology. Anti-human HIF-1α (sc-53546, dilution 1:200), VEGF (sc-152, dilution 1:200), EPO (sc-7956, dilution 1:1 000), total EPOR (sc-697, dilution 1:1 000) and p-EPOR (sc-20236, dilution 1:1 000) antibodies were purchased from Santa Cruz Biotechnology. Equal loading was confirmed with β-actin (AC-15, dilution 1:10 000, Sigma-Aldrich) [17]. Stained proteins were detected using the ECL Plus Western Blotting Detection System (GE Healthcare).

Human renal cells Caki-1, 786-O, 769-P and RPTEC were plated in 96 well dishes in triplicate (10³ cells/well) and incubated in normoxic condition. Cells were then subjected to increasing doses of rhEPO (0–50 units/mL) and incubated in normoxic or hypoxic conditions. After 48 hrs, cell proliferation was determined by CellTiter-Glo Luminescent cell viability assay (Promega, Madison, WI) according to manufacturer’s instructions. Luminescence was measured using a FLUOstar Optima Reader (BMG LABTECH, Ortenberg, Germany). Three independent experiments were performed in triplicate.

Human renal cells were seeded in 6-well plates at a density of 2 × 10⁵ cells per well and incubated for 24 hrs. Cells were starved for 18 hrs in serum/growth factors-free media containing 0.1% BSA in normoxic or hypoxic condition. After starvation, media were replaced with fresh media containing 2% FBS with or without 2 units/mL of rhEPO and incubated for 10 hrs in normoxic or hypoxic condition. Cells were harvested and fixed with 70% ethanol overnight at -20°C. Next, cells were suspended in propidium iodide (PI) staining buffer containing 50 μg/ml PI and 200 μg/ml RNase A and incubated in 37°C for 15 min. PI fluorescence was determined by flow cytometry using a FACSCalibur and CellQuest software for acquisition (BD Biosciences, San Jose, California). Cell cycle phase distribution was analyzed and reported by using FlowJo software (TreeStar Inc., Ashland, OR). Three independent experiments were performed in triplicate.

To obtain populations of cells in G₀/G₁ phase, all human renal cells were arrested by double thymidine block as described previously [18]. Briefly, human renal cells were seeded at 5 × 10⁴ cells per well in a 6-well plate. Cells were blocked for 18 hrs with 2.5 mM thymidine (Sigma-Aldrich), released for 6 hrs, washed to remove the thymidine, and then exposed again to 2.5 mM thymidine this time for 16 hrs in normoxia or hypoxia. The cells were then released from the double thymidine block by culturing in 2% FBS-containing fresh media with or without 2 units/mL of rhEPO and allowed to progress through G1 and into S-phase. The percentage of proliferating cells was determined at 0, 2, 4, 6, 9 and 12 hrs after release from the double thymidine block using the Click-iT® EdU Alexa Fluor® 647 Flow Cytometry Assay Kit (Life Technologies, Carlsbad, CA) according to the manufacturer's instructions. EdU (5-ethynyl-2´-deoxyuridine) is a thymidine analog that becomes incorporated into DNA during active cellular DNA synthesis. Detection is determined via a copper catalyzed covalent reaction between an azide (conjugated to Alexa Fluor 647) and an alkyne. EdU (10 μM) was added to each well 2 hrs prior to harvesting. Cells were trypsinized and fixed in 4% formaldehyde. Cell Quest Pro Software determined cellular DNA synthesis using FlowJo Software. Three independent experiments were performed in triplicate.

Animal care was in compliance with the recommendations of The Guide for Care and Use of Laboratory Animals (National Research Council) and approved by our local IACUC. The subcutaneous tumorigenicity assay was performed in athymic BALB/c (nu/nu) mice, 6 to 8 weeks old purchased from Harlan Laboratories (Indianapolis, IN). Procrit (epoetin α; Amgen Inc, Thousand Oaks, CA) was used for the in vivo treatment of EPO. The properties of rhEPO were tested in vivo using a subcutaneous xenograft model by inoculating 10⁶ Caki-1, 786-O and 769-P cells as described previously [16, 19]. Since RPTEC cells are benign and not known to produce xenograft tumors, this cell line was not tested in vivo. After 24 hrs, mice were divided randomly into two groups (control or 200 international units (IU)/kg of rhEPO) and treatment was initiated. RhEPO was administered subcutaneously once weekly. Control mice received vehicle alone (PBS) on the same schedule. At least 10 animals were in each group. Tumor volumes were measured twice weekly with digital calipers and calculated by V (mm³) = length × (width)² × 0.5236. After 10 wks of treatment, the mice were sacrificed. However, 30 mins before being sacrificed, each mouse was intraperitoneally injected with 0.1 mL (60 mg/kg of body weight) of pimonidazole hydrochloride (Hypoxyprobe-1 Plus Kit; Hypoxyprobe Inc., Burlington, MA), according to the manufacturer's instructions [20]. Subsequently, the mice were sacrificed and xenografts resected. The excised tumors were placed in 10% buffered formaldehyde solution and embedded in paraffin. Paraffin blocks were sectioned for H&E staining and immunohistochemical (IHC) staining.

Paraffin embedded tumors were sectioned (4 μm), deparaffinized in xylene and rehydrated using graded percentages of ethanol. Slides were treated with 1% hydrogen peroxide in methanol to block endogenous peroxidase activity. Staining was conducted using anti-human EPO antibody (sc-7956, dilution 1:200), anti-human EPOR antibody (sc-695, dilution 1:100), HIF-1α (sc-53546, dilution 1:100), VEGF (sc-152, dilution 1:200), cyclin D1 (#2978, dilution 1:50), p21^cip1 (#2947, dilution 1:100), p27^kip1 (#3686, dilution 1:200), anti-human Ki-67 (MIB-1, dilution, 1:200; Dako). Biotin-labeled horse anti-mouse IgG or rabbit IgG (2 μg/ml in 1% BSA blocking buffer) was used as secondary antibody. Immunoreactive signals were amplified by formation of avidin-biotin peroxidase complexes and visualized using 3, 3'- diaminobenzidine (DAB). Nuclear counterstaining was conducted with hematoxylin. Proliferative index analysis was determined as previously described [16]. In addition, slides were immunostained with fluorescein isothiocyanate (FITC)–conjugated primary antibody against pimonidazole (1:50) and horseradish peroxidase–labeled secondary anti-FITC monoclonal antibody (1:50) supplied with the hypoxia detection kit (Hypoxyprobe-1 Plus Kit), according to a modification of the manufacturer's instructions as described previously [20].

All data are expressed as mean ± standard deviation (SD) and mean ± standard error of the mean (SEM). Statistical analyses were conducted using GraphPad Prism 5.0 (GraphPad Software, Inc.). The comparison between EPO and EPOR expression in cancer vs. benign tissue was calculated using Mann–Whitney U test. For most in vitro and in vivo comparisons, a 2-tailed unpaired Student t test or Mann–Whitney U test was conducted. Differences were considered statistically significant at p < 0.05.

We analyzed a human cancer TMA consisting of malignant and benign tissue from 20 organ sites. The immunohistochemistry expression scores from cancerous tissue were compared to those of corresponding benign tissue (Figure 1A). The EPO expression score was significantly elevated in lung cancer (p = 0.003) and lymphoma (p = 0.018). Of note, EPO expression scores in RCC (1.15) and benign renal tissue (1.20) were not significantly different (p = 0.91). Figure 1B shows representative images of EPO immunostaining in lung cancer, lymphoma and RCC. We also scored EPOR expression in the TMA specimens (Figure 1C). The EPOR expression score was significantly elevated in lung (p = 0.011), lymphoma (p = 0.007), thyroid (p = 0.032), uterine (p = 0.038), and prostate cancers (p = 0.011). EPOR expression scores in RCC (1.4) and benign renal tissue (2.0) were not significantly different (p = 0.17). Figure 1D shows representative images of EPOR immunostaining in lung cancer, lymphoma and RCC. The lack of EPO or EPOR correlation to RCC substantiates the previous report by Papworth et al.[21].

We next investigated whether rhEPO might influence cellular proliferation in a panel of human renal cell lines. Key molecules associated with clear cell RCC, as well as EPO and EPOR status were determined in a panel of human renal cell lines comprised of RPTEC, Caki-1, 786-O and 769-P (Figure 2A). We know that expression of the EPO gene is regulated by hypoxia through transcriptional regulators family of hypoxia-inducible factors (HIF) [22], so we also assessed the same key molecules in the cell line panel after exposure to hypoxia over the course of 24 hrs. Hypoxia treatment resulted in the increase of HIF-1α, HIF-2α, EPO and VEGF in all cell lines tested (Figure 2B). A slight increase in EPOR expression was noted in 786-O and 769-P cells exposed to hypoxia, but no changes in VHL expression were observed. We then investigated whether exposing human renal cells to increasing doses of rhEPO could affect cellular proliferation. In an in vitro proliferation assay at 48 hrs, proliferation of RPTEC and Caki-1 cells was significantly enhanced by exposure to 0.5 units/mL rhEPO (p = 0.001) and 2 units/mL rhEPO (p = 0.04), respectively, while the cell lines 786-O and 769-P were unaffected, even at the highest concentration of rhEPO (50 units/mL). Parallel in vitro proliferation assays under hypoxic conditions were also performed. The observed proliferation of RPTEC and Caki-1 cells was significantly enhanced by the exposure of 0.5 units/mL rhEPO (p = 0.0009) and 2 units/mL rhEPO (p = 0.03), respectively. Furthermore, in this hypoxic state, the proliferation of 786-O and 769-P was also significantly increased by the addition of 2 units/mL rhEPO (p = 0.03 and p = 0.04, respectively) (Figure 2C). Thus, in cells with non-functional, mutated VHL (786-O and 769-P) and thus constitutive expression of HIF, rhEPO was able to stimulate cellular proliferation only under hypoxic conditions. Conversely, in cells with functional, wild-type VHL and no HIF expression (RPTEC and Caki-1), rhEPO could stimulate proliferation in both normoxic and hypoxic states.

Standard FACS cell cycle analysis of the panel of cell lines treated with and without rhEPO under normoxic and hypoxic conditions revealed only subtle changes (e.g., S-phase accumulation in RPTEC and 769-P cells treated with rhEPO in hypoxia) (Figure 3). Using a double thymidine block protocol that effectively arrested 98% of the cells at the G₀/G₁-phase of the cell cycle, we were able to more thoroughly assess whether EPO is required for S-phase progression. Cells were released from the double thymidine block by exposing the cells to 2% FBS-containing media with or without 2 units/mL of rhEPO under normoxia or hypoxia (Figure 4A). Synchronized cells of all cell types were more sensitive to rhEPO under hypoxia compared with normoxia. This was more pronounced in RPTEC and 769-P cells. Thus, exposure to rhEPO in a hypoxic state selectively promotes progression from G1 to S-phase, a phase disproportionately represented in frequently dividing cells such as cancer cells. This is the first mention of this phenomenon in the literature.

The expression of molecules that regulate passage of cells from G₀/G₁ to S-phase was analyzed by Western blot (Figure 4B). No significant changes in these molecules were noted in cells exposed to hypoxia, except that p27 ^kip1 was disproportionately elevated relative to cyclin D1 in RPTEC cells. However, upon stimulation with rhEPO in the hypoxic state, cellular levels of cyclin D1 were increased, while cellular levels of p21^cip1 and p27^kip1 were reduced. Conversely, when only rhEPO stimulation was present, only cyclin D1 was increased in RPTEC and Caki-1, and p21^cip1 and p27 ^kip1 were decreased in Caki-1 and 769-P. Our data suggests that in the presence of hypoxia, rhEPO stimulates cellular proliferation in renal cells by promoting progression through G1 into S-phase through upregulation of cyclin D1 and reduction of cell cycle inhibitors (p21^Cip1 and p27^kip1).

Previous studies have linked EPO-induced changes to activation of JAK2 and MAPK-ERK1/2 pathways in some model systems [23‐26]. To confirm that the proliferative effects of EPO are mediated through the activation of JAK2 and MAPK-ERK1/2 in human renal cells, and to evaluate if these same pathways are involved when cells are subjected to a hypoxic environment, we monitored the expression of JAK2, phosphorylated JAK2 (p-JAK2), Stat5 and phosphorylated Stat5 (p-Stat5) to assess the JAK2 pathway, and Akt, phosphorylated Akt (p-Akt), ERK1/2 and phosphorylated ERK1/2 (p-ERK1/2) to assess the MAPK-ERK1/2 pathway. Under normoxic conditions, exposure to rhEPO resulted in an increase in the expression of p-JAK2 and p-ERK1/2 in RPTEC cells, an increase in p-JAK2 in Caki-1 cells, and an increase in p-JAK2, p-AKT and p-ERK1/2 in 786-O cells. No changes were observed in 769-P cells (Figure 5A). Hypoxic culture alone was associated with an increase in the expression of p-ERK1/2 in RPTEC cells, p-JAK2 in Caki-1 cells, p-JAK2 in 786-O and p-JAK2 and p-Akt in 769-P cells. Most notably, in the hypoxic state, the addition of EPO consistently increased the expression of p-JAK2 and p-ERK1/2 in all four cell lines (Figure 5A). Subsequently, we set out to evaluate which pathway, JAK2 or MAPK-ERK1/2, was involved in the observed molecular changes associated with G1-phase progression. This was achieved by targeting each pathway with a small molecule inhibitor (TG101348 targets JAK2 and U0126 targets MEK in the MAPK-ERK1/2 pathway). In all cell lines, and under all experimental conditions (+/− rhEPO and hypoxia/normoxia), TG101348 treatment resulted in a reduction in p-JAK2, and U0126 treatment resulted in a reduction of p-ERK1/2 (Figure 5B). In parallel experiments utilizing these inhibitors, we assessed changes in cell proliferation (Additional file 1: Figure S1), specifically G1-phase progression by Western blot analysis, which documented changes in cyclin D1, p21^cip1 and p27^kip1 expression (Figure 5C). We conclude that EPO exposure results in the activation of both the JAK2 and ERK1/2 pathways leading to changes in proliferation under hypoxic conditions.

To determine whether EPO can regulate tumor growth and proliferation in vivo, we injected subcutaneously Caki-1, 786-O and 769-P cells in athymic nude mice, however, 769-P cells did not form subcutaneous tumors in this model. Systemic administration of rhEPO over the experimental term of 10 wks resulted in a remarkable increase in 786-O tumor size compared to control. Specifically, at the end of the study, control 786-O xenografts achieved an average volume of 603 mm³ compared to 1107 mm³ (p = 0.015) for 786-O tumors treated with 200 IU/mg/week (Figure 6A). However, administration of EPO in Caki-1 xenografts did not result in a tumor growth advantage compared to controls (p = 0.20) (Figure 6A). Evaluation of excised xenografts revealed a clear increase in cyclin D1 and a reduction in p21^cip1 and p27^kip1 in EPO-treated 786-O tumors (Figure 6B). Furthermore, an increase in p-EPOR expression was noted in 786-O xenograft tumors compared to 786-O xenograft controls (Figure 6B). Immunostaining of Caki-1 xenograft tumors are depicted in Additional file 2: Figure S2. The proliferative marker, Ki-67, was studied within the tumor sections and an enhanced Ki-67 positivity was noted in EPO-treated 786-O xenograft tumors. No changes in proliferative index were noted in Caki-1 xenografts treated with rhEPO (Figure 6C). Our in vitro data suggested that hypoxia potentiates rhEPO proliferative effects. So at the termination of the in vivo experiment, pimonidazole staining assessed the extent of xenograft hypoxia. Interestingly, in the Caki-1 xenografts, which had no increase in tumor growth when exposed to rhEPO, limited areas of hypoxia were noted. Conversely, the 786-O xenografts had a considerable number of hypoxic regions (Figure 6D). These in vivo observations confirm the potential of EPO to stimulate cellular proliferation and, hence, tumor growth, especially in a hypoxic setting.