We used several human RMS cell lines (provided by Dr Peter Houghton, Nationwide Children's Cancer Center, Columbus, OH, USA), including both fusion-positive (RH28, RH30 and RH41) and fusion-negative (JR, RD, RH18, RH36 and SMS-CTR) cell lines. All cell lines used in these studies were authenticated by short tandem repeat (STR) analysis. STR profiles were compared with those of the original cell lines, obtained in Dr Peter Houghton's laboratory, or with published profiles. SMS-CTR and RH36 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS), 100 U/ml penicillin and 10 μg/ml streptomycin. All other cell lines were maintained in Roswell Park Memorial Institute (RPMI)-1640 medium, containing 10% FBS, 100 U/ml penicillin and 10 μg/ml streptomycin. All cells were cultured in a humidified atmosphere of 5% CO₂ at 37°C, and the medium was changed every 48 h.

Total RNA was isolated from RMS cells with the RNeasy kit (Qiagen, Inc., Valencia, CA, USA). Human muscle RNA was obtained from Ambion (Austin, TX, USA). The RNA was reverse-transcribed with MultiScribe Reverse Transcriptase, oligo(dT) and a random hexamer primer mix (all from Applied Biosystems Life Technologies, Foster City, CA, USA). Quantitative assessment of the mRNA levels of target genes was performed by qRT-PCR using an ABI Prism 7500 sequence detection system (Applied Biosystems Life Technologies) with Power SYBR-Green PCR Master mix reagent and specific primers (hEpoR forward, CCA TGG ACA CTG TGC CCT G and reverse, CCA TCG GAT AAG CCC CCT T; hEPO forward, CAC CAC GCC TCA TCT GTG AC and reverse, CAC AGC CCG TCG TGA TAT TCT). The PCR cycling conditions were as follows: 95°C (15 sec), 40 cycles at 95°C (15 sec) and 60°C (1 min). According to melting point analysis, only one PCR product was amplified under these conditions. The relative quantity of a target, normalized to the endogenous β2-microglobulin gene as control and relative to a calibrator, is expressed as 2^−ΔΔCt (fold difference), where ΔCt is the threshold cycle, Ct = (Ct of target genes)-(Ct of the endogenous control gene, β2-microglobulin), and ΔΔCt = (ΔCt for target gene in test sample)-(ΔCt for target gene in calibrator sample).

Total RNA from various cells was isolated using the RNeasy Mini kit, including treatment with DNase I (both from Qiagen, Inc.). The mRNA (200 ng) was reverse-transcribed with TaqMan Reverse Transcription Reagents (Applied Biosystems Life Technologies) according to the manufacturer's instructions. The resulting cDNA fragments were amplified (1 cycle of 8 min at 95°C, 2 cycles of 2 min at 95°C, 1 min at 60°C, 1 min at 72°C, and subsequently 35 cycles of 30 sec at 95°C, 1 min at 60°C, 1 min at 72°C, and 1 cycle of 10 min at 72°C) using AmpliTaq Gold with sequence-specific primers designed using the NCBI/Primer-BLAST program; one primer was designed to include an exon-intron boundary (β-actin forward, GGA TGC AGA AGG AGA TCA CTG and reverse, CGA TCC ACA CGG AGT ACT TG; hEpoR forward, CTC CCT TTG TCT CCT GCT CG and reverse, TAG GCA GCG AAC ACC AGA AG; hEPO forward, TCA TCT GTG ACA GCC GAG TC and reverse, GCC ACT GAC GGC TTT ATC CA; and hCD131 forward, AGG GGC AGA AGA AAC CAT CC and reverse, GGC CTG TCT GGT TGG AAT GA).

Cells were grown in 24-well culture plates at an initial density of 1.5×10⁴ cells/well. After 24 h, the medium was changed to new medium supplemented with 10% FBS, and the cells were cultured in the presence or absence of VCR (1 nM to 1 μM). The complete medium (with 10% FBS) was used as a control. The cell number was calculated at 24, 48 and 72 h after the change of medium. At the indicated time points, cells were harvested from the culture plates by trypsinization.

Chemotaxis assays were performed in a modified Boyden's chamber with 8-μm pore polycarbonate membrane inserts (Costar Transwell; Corning Costar, Lowell, MA, USA) as previously described (20). In brief, cells detached with 0.25% trypsin were seeded into the upper chamber of an insert at a density of 4.5×10⁴ in 100 μl. The lower chamber was filled with pre-warmed culture medium containing test reagents. Medium supplemented with 0.5% BSA was used as a negative control. After 24 h, the inserts were removed from the Transwell supports. The cells that had not migrated were scraped off with cotton wool from the upper membrane, and the cells that had transmigrated to the lower side of the membrane were fixed and stained with HEMA 3 (protocol; Thermo Fisher Scientific, Pittsburgh, PA, USA) and counted on the lower side of the membrane using an inverted microscope.

A hypoxic condition was achieved by using a nitrogen-balanced hypoxia chamber (BioSpherix Ltd., Lacona, NY, USA) providing a gas mixture containing 5% CO₂ and 2% O₂ at 37°C.

RMS cell lines were incubated overnight in RPMI-1640 medium containing low levels of bovine serum albumin (BSA, 0.5%) to render the cells quiescent. After the cells were stimulated with EPO (0.5 and 20 IU/ml) at 37°C for 5 min, the cells were lysed for 10 min on ice in RIPA lysis buffer containing protease and phosphatase inhibitors (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA). The extracted proteins were separated on a 4–12% SDS-PAGE gel and transferred to a PVDF membrane. Phosphorylation of the serine/threonine kinase AKT (phospho-AKT473) and p44/42 mitogen-activated kinase (phospho-p44/42 MAPK) was detected by phospho-specific p44/42 MAPK mouse and rabbit polyclonal antibodies (Cell Signaling Technology, Inc., Danvers, MA, USA) with HRP-conjugated goat anti-mouse and anti-rabbit secondary antibodies (Santa Cruz Biotechnology, Inc.). Equal loading in the lanes was evaluated by stripping the blots and reprobing with anti-p42/44 MAPK monoclonal antibody (clone no. 9102) and anti-AKT polyclonal antibody (both from Cell Signaling Technology, Inc.). The membranes were developed with an enhanced chemiluminescence (ECL) reagent (Amersham Life Sciences, Arlington Heights, IL, USA), dried, and subsequently exposed to film (Hyperfilm; Amersham Life Sciences).

Cells were made quiescent for 3 h with 0.5% BSA in RPMI-1640 before incubation with EPO (2 and 20 IU/ml). Subsequently, cell suspensions (2×10³/100 μl) were added directly to 96-well plates covered with fibronectin and incubated for 5 min at 37°C. The wells were coated with fibronectin (10 μg/ml) overnight at 4°C and blocked with 0.5% BSA for 1 h before the experiment. Following incubation, the plates were vigorously washed three times to remove non-adherent cells, and the adherent cells were counted using an inverted microscope.

To evaluate the effect of hypoxia on expression of CD131 and endogenous EPO and EpoR in RMS cells, cells were incubated in hypoxic condition by using a nitrogen-balanced hypoxia chamber providing a gas mixture containing 5% CO₂ and 2% O₂ at 37°C. Cells were treated for 24 h and subsequently RNA has been extracted for RT-PCR analysis using RNeasy kit (Qiagen, Inc.).

To evaluate expression of EpoR in fusion positive and negative RMS cells, the cells were incubated with human EpoR antibody (APC, clone 38421) for 30 min on ice. After washing, cells were evaluated using a BD LSR II flow cytometer (BD Biosciences, San Jose, CA, USA). Representative FACS results are shown. FACS analysis was also employed in proliferation assay in the presence of VCR (1 nM-1 μM) or EPO (20 IU/ml) and in apoptosis assay employing Annexin V-PE.

RMS cells were cultured with or without EPO (20 IU/ml) and with or without VCR (1 μM) in serum-free medium for 24, 48 and 72 h. Apoptosis was detected by FACS using the Annexin V-PE Apoptosis Detection kit (BD Pharmingen, San Diego, CA, USA). Cells were stained for apoptotic cell detection with Annexin V-PE for 15 min at RT.

Fifty-eight RMS frozen primary tumor specimens (26 PAX3-FOXO1-positive, 7 PAX7-FOXO1-positive, and 25 fusion-negative) were used for microarray profiling. Total RNA was extracted from primary tumors using RNA STAT-60 (Tel-Test, Friendswood, TX, USA) in accordance with manufacturer's instructions. These RNA samples were analyzed on Affymetrix GeneChip^® Human Genome U133 Plus 2.0 Arrays at the University of Pennsylvania Microarray Core Facility. The expression array data were normalized and log2 transformed with Robus Multiarray Average (RMA) in the Bioconductor oligo package. The expression level of EPOR was calculated as an average of three probe sets (probe set ID: 215054_at, 209962_at and 37986_at). Probe set ID 215059_at is the only available probe set for CD131 on the U133 Plus 2.0 array.

The results are presented as mean ± SD. Statistical analysis of the data was done using Student's t-test for unpaired samples, with P<0.05 considered significant.

First, we performed RT-PCR studies to evaluate mRNA expression in three human fusion-positive (RH28, RH30 and RH41) as well as five human fusion-negative (JR, RD, RH18, RH36 and SMS-CTR) RMS cell lines. Fig. 1A, shows that we were able to detect EpoR mRNA by regular RT-PCR in all cell lines employed in this study, but not in skeletal muscle cells. As shown in Fig. 1B, all cell lines, especially RH18 and RH36, expressed high levels of EpoR mRNA as determined by quantitative RT-PCR. Of note very week signal was also detected in skeletal muscle cells by employing this highly sensitive cDNA amplification method. Expression of EpoR was subsequently confirmed by FACS (Fig. 1C), and RH36 and RH18 highly expressed this receptor at the protein level. No expression was detected on skeletal muscle cells (data not shown).

Since it is known that EPO signals via EpoR-EpoR homodimers in erythroid cells and may signal via EpoR-CD131 heterodimers in non-erythroid cells (21,22), we evaluated the expression of CD131 in our RMS cell lines by FACS and RT-PCR, and found that CD131 was not detectably expressed in these cells (data not shown). However, when we exposed RMS cells to hypoxia, CD133 become detectable (Fig. 2A). In parallel we observed upregulation of EpoR and EpO (Fig. 2B). Furthermore, since our RT-PCR analysis revealed expression of endogenous EpO in studied RMS cell lines (Fig. 3), we cannot exclude contribution of autocrine EpO-EpoR interactions, and this phenomenon is currently investigated in our laboratory.

Next, to evaluate whether EpoR is functional in human RMS cell lines, we performed chemotaxis and cell adhesion studies. In Transwell chemotaxis assays (Fig. 4A) we employed EPO at doses of 5, 20 and 50 U/ml as chemoattractant and compared the responsiveness of RMS cells to 300 ng/ml of the α-chemokine stromal-derived factor 1 (SDF-1), which, as we previously described (23,24), is a known chemoattractant for these cells. We observed that EPO induced motility of all RMS cell lines evaluated in our studies. The highest responsiveness was observed for RH30 (fusion-positive) cell lines and for RH18, RH30, RH36 and RH28 cell lines. The specificity of EpO-EpoR interaction has been confirmed by employing EpoR blocking antibodies (data not shown).

The motility of cancer cells plays a crucial role in the process of tumor metastasis, and to address whether the observed increase in motility of RMS cells in response to EPO is the result of a chemotactic or a chemokinetic response, we performed a checkerboard assay in which EPO was added at the same time into the upper and the lower Transwell chambers so that there was no EPO gradient between the chambers. Fig. 4B, demonstrates that the migration of RH30 cells in response to EPO in the upper and lower chambers was not significantly changed when it was in both chambers compared to only one, the lower chamber. This finding indicates that EPO is mainly a chemokinetic rather than a chemotactic factor for RMS cells.

Another important feature of metastasizing cancer cells is their adhesive properties at the site of metastasis. Therefore, we next studied the effect of EPO on adhesion of RMS cell lines to fibronectin. Depending on the dose employed, we found that EPO may induce adhesion of RMS cells to fibronectin (Fig. 4C), and this effect was particularly visible for RH28 and SMS-CTR cells. Again, the specificity of EpO-EpoR interaction was confirmed by employing EpoR blocking antibodies (data not shown).

EPO is a known growth factor for erythroid cells (22) as well as for some tumor cell lines (5–14). Therefore, based on receptor expression studies (Fig. 1), we exposed RH18 cell line and the RH36 cell line to 20 IU/ml EPO and evaluated its effect on proliferation of these cells. We found that EPO stimulated proliferation of both RH18 (Fig. 5A) and RH36 cells (Fig. 5B). Western blot analysis confirmed that both cell lines responded to EPO stimulation by phosphorylation of MAPK^p44/42 and to a lesser degree also by phosphorylation of Akt^ser473.

Since EPO has pro-proliferative effects on RMS cells, to assess its effect on the potential resistance of RMS cells to chemotherapy, we performed proliferation assays of these cells in medium supplemented with EPO and different doses of VCR. As demonstrated in Fig. 6, we observed an increase in survival of RH18 (Fig. 6A) and RH36 cells (Fig. 6B) in the presence of VCR (25). While a significant beneficial effect was observed for RH18 cells after and 72 h of treatment, some beneficial effect for RH36 cells was observed after only 24 h of treatment. The cell proliferation data are supported by the effect of EpO on inhibiting apoptosis in VCR treated cells (data not shown).

Finally, we assessed EpoR expression from expression microarray analyses of 26 PAX3-FOXO1-positive, 7 PAX7-FOXO1-positive, and 25 fusion-negative patient samples. In all these tumor samples, we were able to detect EpoR expression; however, expression of EpoR was higher in PAX3-FOXO1-positive and in fusion-negative samples than in PAX7-FOXO1-positive samples (Fig. 7A). Interestingly, in contrast to established RMS cell lines (data not shown), CD131 expression was detected in all human RMS samples (Fig. 7B), and similar to our EpoR findings, expression of CD133 was higher in PAX3-FOXO1-positive as compared to PAX7-FOXO1-positive tumor samples.