Advertisement
Angiogenesis, Metastasis, and the Cellular Microenvironment

Regulation of Expression of Stromal-Derived Factor-1 Receptors: CXCR4 and CXCR7 in Human Rhabdomyosarcomas

Maciej Tarnowski, Katarzyna Grymula, Ryan Reca, Kacper Jankowski, Radoslaw Maksym, Joanna Tarnowska, Grzegorz Przybylski, Frederic G. Barr, Magdalena Kucia and Mariusz Z. Ratajczak
DOI: 10.1158/1541-7786.MCR-09-0259 Published January 2010

Abstract

Rhabdomyosarcomas (RMS) express CXCR4 and CXCR7 receptors that bind prometastatic α-chemokine stromal-derived factor-1 (SDF-1). In this report, we analyzed the activity of both promoters in a model of less metastatic human embryonal-RMS cell line (RD) and more metastatic alveolar-like RMS (RD cells transduced with paired box gene 3/forkhead homologue; PAX3-FKHR fusion gene). First, CXCR4 is barely detectable in RD and becomes upregulated in RD/PAX3-FKHR cells. In contrast, CXCR7 highly expressed in RD becomes downregulated in RD/PAX3-FKHR cells. Next, promoter deletion and mutation studies revealed that whereas (a) expression of CXCR4 in RD and RD/PAX3-FKHR cells required nuclear respiratory factor-1 (NRF-1) binding site and (b) was additionally upregulated by direct interaction of NRF-1 with PAX3-FKHR, CXCR7 promoter activity required a proximal nuclear factor-κB–binding motif. The requirement of these factors for CXCR4 and CXCR7 promoter activities was additionally supported after blocking NRF-1 and nuclear factor-κB. Furthermore, CXCR4 expression in PAX3-FKHR+ RMS cells seems to be enhanced because of the interaction of PAX3-FKHR and NRF-1 proteins in the proximal part of the promoter that prevents access of the negative regulator of transcription YY1 to its binding site. Finally, although hypoxia enhances CXCR4 and CXCR7 promoter activity and receptor expression in RD cells, it inhibits CXCR7 expression in RD/PAX3-FKHR cells. In conclusion, SDF-1 binding receptors CXCR4 and CXCR7 are differently regulated in RMS cells. The upregulation of CXCR4 and downregulation of CXCR7 expression by PAX3-FKHR or hypoxia may give SDF-1 an advantage to better engage the CXCR4 receptor, thus increasing RMS motility. Mol Cancer Res; 8(1); 1–14

Keywords
  • Rhabdomyosarcoma
  • SDF-1
  • CXCR4
  • CXCR7
  • PAX3-FKHR

Introduction

Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma of adolescence and childhood and accounts for 5% of all malignant tumors in patients under 15 years of age (1-10). There are two major histologic subtypes of RMS, alveolar (A)RMS and embryonal (E)RMS. Clinical evidence indicates that ARMS is more aggressive and has a significantly worse outcome than ERMS (11, 12). Genetic characterization of RMS has identified markers that show excellent correlation with histologic subtype. Specifically, ARMS is characterized by the translocation t(2;13)(q35;q14) in 70% of cases or the variant t(1;13)(p36;q14) in a smaller percentage of cases. These translocations disrupt the paired box genes (PAX3 and PAX7) on chromosomes 2 and 1, respectively, and the forkhead homologue in RMS (FKHR) gene on chromosome 13 and generate PAX3-FKHR and PAX7-FKHR fusion genes. These fusion genes encode the fusion proteins PAX3-FKHR and PAX7-FKHR, which increase the transcription factor activity of PAX3/7 proteins and enhance the metastatic potential of ARMS cells (13).

Chemokines, the small proinflammatory chemoattractant cytokines that bind to specific G protein–coupled, seven transmembrane receptors present on the plasma membranes of target cells, are the major regulators of trafficking and adhesion of normal and malignant cells (14, 15). More than 50 different chemokines have been cloned thus far and some of them were reported to play a pivotal role in cancer metastasis. One of the most important is α-chemokine stromal-derived factor-1 (SDF-1), which regulates the metastatic behavior of several types of cancer (16-19). SDF-1 binds to G protein–coupled, seven transmembrane–span receptor, CXCR4. In our previous work, we showed a pivotal role of the SDF-1-CXCR4 axis in the metastasis of human RMS to various organs including bone marrow (17).

For many years, it was postulated that CXCR4 was the only receptor for SDF-1. However, the concept of an exclusive interaction of SDF-1 with CXCR4 was questioned recently after a new SDF-1 binding receptor, CXCR7, was identified (20). CXCR7, in contrast to CXCR4, has another ligand, a chemokine called IFN-inducible T-cell chemoattractant (I-TAC; ref. 20). Recently, we noticed that the SDF-1-CXCR7 axis regulates the metastatic potential of human RMS cells similarly to SDF-1-CXCR4 (21). We observed that whereas SDF-1 signaling through CXCR4 enhances more significant RMS cell motility, CXCR7 is somehow more involved in the adhesion of RMS cells. Both these receptors, however, are potential targets for new antimetastatic strategies (22-24).

To learn more about the SDF-1-CXCR4 and SDF-1-CXCR7 axes in RMS metastasis, we analyzed the activity of both promoters in a model of less metastatic embryonal RD cells and more metastatic (alveolar-like) RD cells transduced with PAX3-FKHR fusion gene (RD/PAX3-FKHR). By using promoter deletion and mutation studies, we found that CXCR4 promoter activity depends on a proximal nuclear respiratory factor-1 (NRF-1) binding site and is enhanced after direct interaction of NRF-1 with PAX3-FKHR. In addition, promoter deletion/mutation and chromatin immunoprecipitation (ChIP) assay studies revealed that NRF-1-PAX3-FKHR interaction in the proximal part of the promoter might prevent access of YY1, which is a negative regulator of transcription to its binding motif located between NRF-1 and proximal PAX3-FKHR binding site. In contrast, CXCR7 promoter activity depends on a proximal nuclear factor-κB (NF-κB) binding site. The requirement of NRF-1 and NF-κB for CXCR4 and CXCR7 promoter activity, respectively, was additionally supported after blocking the expression of these transcription factors by using NRF-1 short hairpin RNA (shRNA) or BAY 11-7082 (small molecular NF-κB inhibitor). We also noticed that CXCR4 and CXCR7 are differently regulated in human RMS cells in response to hypoxia. We postulate that the hypoxia and PAX3/FKHR-mediated upregulation of CXCR4 and downregulation of CXCR7 may provide an advantage for SDF-1 to better engage the CXCR4 receptor. As a result of this, improved SDF-1 interaction with the CXCR4 receptor increases RMS motility and enhances the metastatic potential of RMS cells.

Materials and Methods

Cell Lines and Normoxia Culture Conditions

Three RMS cell lines were used in the experiments: RH30 (a gift from Dr. Peter Houghton, St. Jude Children's Research Hospital, Memphis, TN), RD, and RD-transfected cell lines. All cell lines were maintained in a humidified atmosphere at 5% CO2, 37°C at an initial cell density of 2.5 × 104 cells/flask (Corning) and the medium was changed every 48 h. The RMS cell line (RD) was transfected with empty vector or PAX3-Forkhead box O1 (FOXO1; RD/PAX3-FKHR; ref. 25). Cell cultures were grown in cultured RPMI 1640 (Sigma) supplemented with 100 IU/mL of penicillin, 10 μg/mL of streptomycin, and 50 μg/mL of neomycin (Life Technologies, Inc.) in the presence of 10% heat-inactivated fetal bovine serum (Life Technologies) and 300 μg/mL of G-418.

Cell Surface Expression of CXCR4 and CXCR7

Cell surface expression was measured by flow cytometry. Cells were stained for surface CXCR4 with APC-conjugated antibody anti-CXCR4 (clone no. 12G5; BD Biosciences) and for surface CXCR7 with phycoerytherin-conjugated antibodies anti-CXCR7 (clone no. 11G8; R&D Systems). Isotype-matched APC and phycoerytherin-conjugated immunoglobulin served as controls (BD Biosciences). All analyses were done on an LSR cell cytometer (BD Biosciences). Samples were evaluated in triplicate and the data were averaged for statistical analysis.

Cloning of the CXCR4 and CXCR7 Promoters

The promoter region of the CXCR4 gene from −2,237 to +62 and the CXCR7 promoter region from −2409 to +89 relative to the transcription start site were amplified. The confirmed sequence was then inserted into a pGL4.10 vector (Promega). The pGL4.10 vector was digested by SacI and XhoI (or KpnI/EcoRV for CXCR7; all restriction enzymes from New England Biolabs) and the amplified CXCR4 and CXCR7 promoter fragments were inserted through ligation. The cloned pGL4.10-CXCR4/7 constructs were confirmed by sequencing. The sequentially shorter CXCR4/7 promoter fragments were amplified by standard PCR methods, sequenced, and cloned to pGL4.10 vector. In both CXCR4 and CXCR7 promoters, three mutated constructs were prepared, i.e., CXCR4 NRF-1mut, CXCR4 hypoxia-responsive element (HREmut), and CXCR4 YY1mut as well as CXCR7 NF-κBmut, CXCR7 HREmut, and CXCR7 YY1mut with a QuickChange site-directed mutagenesis kit (Stratagene).

Transient Transfection Assay

RD and RD/PAX3-FKHR cells were transfected with 1.8 μg of CXCR4/CXCR7 constructs and pGL4.72 vector (ratio 50:1) using LipofectAMINE (Invitrogen) in 12-well plates according to the protocols of the manufacturer. At 24 h after transfection, cells were lysed and 10 μL of each sample was analyzed for firefly/Renilla luciferase activity with the dual luciferase assay system (Promega) and measured on a luminometer (Turner Biosystems) with the software provided. The firefly/Renilla luciferase activity ratios were calculated with that software and used to evaluate the changes in promoter activity in hypoxia and normoxia. Fold difference was based on empty pGL4.10 vector activity.

Immunoprecipitation and Western Blot

The 5 × 106 cells were lysed in 1 mL of lysis buffer [50 mmol/L Tris-HCl (pH 8.0), 150 mmol/L NaCl, 1% Triton X-100, and protease inhibitors] for 10 min on ice, syringed eight times with a 21-gauge syringe, and spun down for 15 min at 10,000 × g at 4°C. Supernatants were transferred to new tubes and precleared with 100 μL of Preclearing Matrix (Santa Cruz Biotechnology) rotating overnight at 4°C. Samples were spun down for 1 min at full speed and supernatants were incubated with 0.2 μg/mL of a goat anti-human PAX3/7 polyclonal antibody clone N-19 (Santa Cruz Biotechnology) and IP Matrix F (Santa Cruz Biotechnology) rotating overnight at 4°C. Samples were washed five times with lysis buffer and used for Western blot analysis using a polyclonal rabbit anti-human NRF-1 antibody clone H-285 (Santa Cruz Biotechnology).

ChIP Assay

Either 1 × 106 RD or RD/PAX3-FKHR cells were fixed with 1% formaldehyde (10 min at 37°C), quenched with glycine (12 mmol/L, 5 min at room temperature), lysed, and extracts were used for ChIP analysis according to the protocols of the manufacturer (Upstate Biotechnology). Extracts were sonicated (five times with 10-s pulses with 1-min breaks) on ice with a 60 Sonic Dismembrator (Fisher Scientific). For the CXCR4 promoter, 3 μg of a PAX3/7 antibody clone N-19 or NRF-1 antibody clone H-285 or YY1 clone H-414 (Santa Cruz Biotechnology) and 20 μL of protein G magnetic beads were used to immunoprecipitate protein/DNA complexes. For the CXCR7 promoter, soluble chromatin was incubated with 3 μg of NF-κB P50 antibodies. Negative controls were incubated with rabbit IgG (Santa Cruz Biotechnology). After incubation, complexes were extensively washed and separated on a magnetic separator. Immunoprecipitates were incubated with proteinase K (62°C for 2 h and 95°C for 10 min) and DNA was cleaned up with spin columns and resuspended in Tris-EDTA buffer. Subsequently, a series of PCR reactions were carried out. The sequences of the primers used are shown in Supplementary Table S1.

Real-time Quantitative Reverse Transcription PCR

Total RNA was isolated from cells treated with hypoxia and controls with RNeasy Kit (Qiagen). The RNA was reverse-transcribed with MultiScribe reverse transcriptase and oligo dt primers (Applied Biosystems). Quantitative assessment of mRNA levels was done by real-time quantitative reverse transcription PCR (RQ-PCR) on an ABI 7500 instrument and Power SyBR Green PCR Master Mix reagent. Real-time conditions were as follows: 95°C (15 s), 40 cycles −95°C (15 s), 60°C (1 min). According to a melting point analysis, only one PCR product was amplified under these conditions. The relative quantization value of a target, normalized to the endogenous control β2-microglobulin gene and relative to a calibrator, is expressed as 2-ΔΔCt (fold difference), where ΔCt = (Ct of target genes) − (Ct of endogenous control gene, β2 microglobulin), and ΔΔCt = (ΔCt of samples for target gene) − (ΔCt of calibrator for the target gene). The following primer pairs were used:

  • CXCR4 forward 5-GGTTCCTTCATGGAGTCATAGTC-3,

  • CXCR4 reverse 5-CGGTTACCATGGAGGGGATC-3;

  • CXCR7 forward 5-GGCTATGACACGCACTGCTACA-3,

  • CXCR7 reverse 5-TGGTTGTGCTGCACGAGACT-3;

  • YY1 forward 5-CCCACGGTCCCAGAGTCCA-3,

  • YY1 reverse 5-GTGTGCGCAAATTGAAGTCCAGT-3.

Knockdown of NRF-1 with shRNA and Inhibition of NF-κB with BAY 11-7082

In RNAi experiments, shRNA-generating plasmid pSuper was used. The oligonucleotide targeting base sequence for human NRF-1 was 5′-CATATGGCTACCATAGAAG-3′. Rhabdomyosarcoma cells were plated at 80% confluency and transfected with shRNA vector using LipofectAMINE 2000 (Invitrogen) according to the protocols of the manufacturer. Commercially available, negative-scrambled control plasmid was used (Dharmacon). NF-κB inhibitor BAY 11-7082 was purchased from Sigma. Cells were treated for 16 h with 1 μmol/L of inhibitor and then receptor expression was analyzed as described before. In the hypoxia assay, cells were pretreated for 30 min with the inhibitor and then subjected to hypoxia.

Hypoxia Assay

A hypoxic condition was acquired using a nitrogen-balanced hypoxia chamber providing a gas mixture containing 5% CO2 and 1% O2 at 37°C. In the assay, three RMS cell lines were used (RD, RD/PAX3-FKHR, and RH30). Cells were treated overnight or for different periods of time, and then subjected to fluorescence-activated cell sorting analysis or dual luciferase assay.

Transmembrane Chemotaxis

The 8-μm pore polycarbonate membranes were covered with 50 μL of 0.5% gelatin. Cells were detached with 0.5 mmol/L of EDTA, washed, resuspended in RPMI 1640 with 0.5% bovine serum albumin, and seeded at a density of 3 × 104 in 120 μL into the upper chambers of Transwell inserts (Corning Costar, Corning). The lower chambers were filled with medium alone or medium containing SDF-1 (300 ng/mL) or I-TAC (100 ng/mL), conditioned medium, or 0.5% bovine serum albumin RPMI 1640 (control). Plates were put into normoxic and hypoxic conditions. After 24 h, the inserts were removed from the Transwells. Cells remaining in the upper chambers were scraped off with cotton wool. Cells that had transmigrated were stained by hydroxyethyl methacrylate 3 (Protocol, Fisher Scientific) and counted either on the lower side of the membranes.

Statistical Analysis

All results were presented as mean ± SEM. Statistical analysis of the data was done using the nonparametric Mann-Whitney test and Student's t test for unpaired samples, with P < 0.05 considered significant.

Results

CXCR4 and CXCR7 Are Differently Expressed on RD and RD/PAX3-FKHR Cells

As previously reported, human RMS cells express both SDF-1 binding receptors (17, 26). Figure 1A shows the cell surface expression of CXCR4 and CXCR7 at the protein level on human RD and RD/PAX3-FKHR cell lines used in the current study. RD cells that belong to the ERMS subtype highly express CXCR7 and very little CXCR4. In contrast, RD cells stably transduced with PAX3-FKHR construct (ARMS-like subtype) highly express CXCR4 and downregulate the expression of CXCR7. This reciprocal expression of CXCR4 and CXCR7 was subsequently confirmed by RQ-PCR (Fig. 1B). This pattern of expression of CXCR4 and CXCR7 receptors is consistent with the data obtained on nine other human ERMS and ARMS cell lines (21).

FIGURE 1.
FIGURE 1.

A. CXCR4 and CXCR7 membrane expression in RMS cell lines RD and RD/PAX3-FKHR. Flow cytometry analysis of membrane expression was done on the RD cell line transfected with a PAX3-FKHR hybrid construct named RD/PAX3-FKHR and on the RD cell line transfected with empty vector. Percentage of positive cells was calculated on the basis of adequate isotype control (<1%). MFI ± SD. The experiment was repeated thrice with similar results. A representative study is shown. B. RQ-PCR analysis of CXCR4 and CXCR7 mRNA expression in RD and RD/PAX3-FKHR cell lines. The mRNA expression was measured by real-time PCR. Fold of difference was calculated on the basis of 2ΔCt values, in which CXCR4 expression in RD cells = 1 and CXCR7 expression in RD/PAX3-FKHR = 1.

To learn more about the molecular mechanisms governing the expression of both receptors, we cloned CXCR4 and CXCR7 promoters and analyzed the activity of promoter fragments subcloned in a luciferase reporter gene (pGL4.10 vector) in both RD and RD/PAX3-FKHR cell lines.

Cloning the CXCR4 Promoter and Generating CXCR4 Promoter Fragments

The 2299-bp-long sequence of the CXCR4 promoter from −2,237 to +62 was generated from genomic DNA by PCR (Supplementary Fig. S1). The cloned promoter region was sequenced and scanned for possible PAX3 binding sites. We identified 10 putative PAX3 binding sites located at −558 to −572, −596 to −609, −638 to −658, −709 to −721, −1480 to −1510, −1593 to −1597, −1753 to −1757, −1839 to −1848, −2097 to −2101, and −2199 to −2212 of the cloned CXCR4 promoter sequence (Fig. 2A). We also noticed that the CXCR4 promoter contains four HREs (−852 to −856, −1038 to −1042, −1282 to −1286, and −2019 to −2023), one NF-κB (−213 to −223), and one NRF-1 binding site −37 to 54 bp from the transcription start site (Fig. 2A; Supplementary Fig. S1), in addition to putative PAX3 binding sites. Furthermore, at −303 to −307, we identified a binding site for negative transcription regulatory factor YY1. Based on this, nine luciferase reporter gene constructs were generated containing smaller CXCR4 promoter fragments subcloned into a pGL4.10 vector (Fig. 2A). We also generated three promoter constructs in which NRF-1, one HRE, and one YY1 binding site were mutated (Fig. 2A). The HRE mutation construct shown in Fig. 2A (bottom) was also subsequently used in hypoxia and CXCR4 expression studies.

FIGURE 2.
FIGURE 2.

CXCR4 promoter deletion studies and transcription binding sites analysis. A. Constructed CXCR4 promoter inserts. The promoter region of the CXCR4 gene from −2,237 to +62 relative to the start of transcription was cloned and is described as fragment 1 (Frag 1). Fragment 1 was sequentially shortened according to the positions of PAX3 binding sites, and NRF-1 binding sites as depicted in the theoretical model of the cloned sequences of the promoter. The position of PAX3, NRF-1, HRE, NF-κB, and YY1–binding sites as well as P1 to P3 primer pairs used in the ChIP experiment are shown. B. CXCR4 promoter activity studies. RD and RD/PAX3-FKHR cells were transfected with the appropriate plasmids. Cultured cells were harvested after 24 h and assayed for the amount of luciferase activity. Activity was measured on the basis of firefly/Renillla luciferase activity and then the equimolar fold of difference was counted. Results are expressed relative to a value of 1.0 for cells transfected with pGL4.72 vector and empty pGL4.10. Averages of duplicates from three independent experiments are shown. Values are given as the mean ± SEM. Frag1-7 P < 0.05, as compared with empty vector. *, P < 0.05, fragments compared with NRF-1mut fragments and Frag8 (Mann-Whitney test). C. The PAX3-FKHR-NRF-1 complex binds DNA. The ChIP assay was done on RD/PAX3-FKHR nuclear extracts pulled down with the anti-PAX3/7 antibodies and revealed that two of the PAX3 binding sites assayed were preserved and that the NRF-1 binding site was preserved as well. Moreover, pulling down with NRF antibodies resulted in the preservation of both PAX3 binding sites. In the RD cell line, aside from the NRF binding site, one PAX3 binding site seemed to be occupied by transcription factors. No complex formation was visible in RD cells. A representative example is shown from three independent experiments.

PAX3-FKHR Regulation of the CXCR4 Promoter

Next, RD and RD/PAX3-FKHR cells were transfected by a luciferase reporter gene that was driven by different CXCR4 promoter fragments. Overall, we found a higher luciferase level in RD cells that were transduced with PAX3-FKHR (Fig. 2B). We found that CXCR4 Frag 7, which contains a NRF-1 binding site but not PAX3 domains, is still able to drive the luciferase gene in both RD and even at a much higher level in RD/PAX3-FKHR cells. This could be explained by the 124 bp fragment of the CXCR4 promoter possibly containing some cryptic PAX3 binding site that is responsible for higher activity of the CXCR4 promoter in the latter cells in the presence of PAX3-FKHR fusion protein. Subsequently, deletion of NRF-1 completely abrogated promoter activity as seen in cells transduced with the shortest CXCR4 Frag 8. To confirm that NRF-1 is crucial for the expression of the CXCR4 promoter, RD and RD-PAX3/FKHR cells were transduced with a whole promoter fragment in which an NRF-1 binding site was mutated (NRF-1mut; Fig. 2A). As shown in Fig. 2B, after NRF-1 mutation, CXCR4 promoter activity was abolished in RD and severely reduced in RD-PAX3-FKHR cells. Thus, our data confirm a crucial role of intact NRF-1 in CXCR4 promoter activity (27, 28). They also indicate that PAX3-FKHR, which is a strong transcription factor, may additionally increase CXCR4 promoter activity in RMS cells. Moreover, PAX3-FKHR seems to maintain some level of CXCR4 promoter activity even when NRF-1 was mutated (Fig. 2B).

PAX3-FKHR Protein Interacts with NRF-1

Next, to confirm that a PAX3-FKHR protein could bind directly to the CXCR4 promoter, we performed ChIP analyses using anti-PAX3/7 and anti-NRF antibodies. By using sets of primers (P3, P2, and P1) designed to amplify different promoter regions that contain PAX3 binding sites and primers (N) designed to amplify the NRF-1 binding region (Fig. 2A; Supplementary Table S1), we detected the presence of PAX3 and NRF-1–protected regions in the CXCR4 promoter (Fig. 2C). We found that PAX3/7 protein binds to the CXCR4 promoter region flanked by P1 primers in both RD and RD/PAX3-FKHR cells, and to the region flanked by P2 primers in RD/PAX3-FKHR cells. Protection of the PAX3 binding site flanked by P2 primers in RD/PAX3-FKHR cells, but not RD cells, is likely explained as being a result of enhanced transcriptional activity of PAX3-FKHR fusion protein.

At the same time, we confirmed NRF-1 protein binding to a proximal fragment of the CXCR4 promoter (Fig. 2C). To our surprise, however, NRF-1 protein protected the proximal PAX3 binding region in RD/PAX3-FKHR cells that was flanked by P1 primers and, at the same time, PAX3-FKHR protein protected the NRF-1 binding site flanked by N primers. This suggests a direct interaction of the PAX3-FKHR fusion protein with the NRF-1 transcription factor. This direct interaction between PAX3-FKHR and NRF-1 in RD/PAX3-FKHR was confirmed by Western blot analysis (Fig. 3A and B), in which the PAX3-FKHR-NRF-1 complex was immunoprecipitated with anti-PAX3/7 antibodies and subsequently probed on the gel with antibodies against NRF-1 (Fig. 3C), as well as when reverse immunoprecipitation was done by using anti–NRF-1 antibodies (Fig. 3D).

FIGURE 3.
FIGURE 3.

PAX3-FKHR binds NRF-1. The NRF-1 binding site was shown to be critical for the increase in CXCR4 promoter activity. A. Western blot analysis of RD and RD/PAX3-FKHR nuclear extracts probed with an anti-PAX3/PAX7 antibody (Santa Cruz Biotechnology). We found a band corresponding to wild-type PAX3 at 56 kDa in both cell lines; however, we also found a band at 97 kDa corresponding to PAX3-FKHR fusion protein. B. Western blot analysis of whole RD and RD/PAX3-FKHR extracts probed with an anti–NRF-1 antibody revealed the presence of a band at 68 kDa corresponding to the predicted size of the NRF-1 protein. C. An immunoprecipitation with both RD and RD/PAX3-FKHR whole cell extracts with the PAX3/7 antibody. The blot was subsequently probed with the anti–NRF-1 antibody. We noticed a band at 68 kDa in the blot of RD/PAX3-FKHR, but not in RD immunoprecipitates, corresponding to the size of NRF-1. D. Reverse experiment, immunoprecipitation with NRF-1 and PAX3 detection shows interaction between NRF-1 and fusion protein PAX3-FKHR. IgG represents the anti-rabbit IgG control for both cell lines.

Cloning the CXCR7 Promoter and Generating CXCR7 Promoter Fragments

CXCR7 promoter was cloned by using DNA-specific primers. We found that the CXCR7 promoter contains three potential HRE (−100 to −104, −965 to −969, −1306 to −1310), five NF-κB (−32 to −42, −308 to −318, −1019 to −1029, −1375 to −1379, −2145 to −2155), and four NRF-binding sites (−1030 to −1040, −1468 to −1478, −1980 to −1990, −2085 to −2095) located within 2.5 kb upstream of the transcriptional start site (Fig. 4A; Supplementary Fig. S2). Furthermore, at −702 to −706, we identified a binding site for negative transcription regulatory factor YY1. Subsequently, we generated eight constructs containing smaller CXCR7 promoter fragments and two constructs containing mutated proximal NF-κB and HRE as well as YY1 binding sites that were subcloned into a pGL4.10 vector (Fig. 4A). The proximal HRE mutation construct shown in Fig. 4A (bottom) was also subsequently used in hypoxia and CXCR7 expression studies.

FIGURE 4.
FIGURE 4.

CXCR7 promoter deletion studies and transcription binding sites analysis. A. CXCR7 promoter inserts. The promoter region of the CXCR7 gene from −2,409 to +89 relative to the start of transcription was cloned and is described as fragment 1 (Frag 1). Fragment 1's shorter derivatives were obtained according to the positions of NRF-1, HRE binding sites, and NF-κB binding sites. The individual constructs were cloned to pGL4.10 vector, transfected into either RD or RD/PAX3-FKHR cells, and used in dual luciferase assays. ×, mutated NF-κB and HRE binding sites. Primer pairs used in the ChIP experiment (N1-N5) flanked five different potential binding sites for NF-κB transcription factor. B. NF-κB as a crucial transcription factor that drives CXCR7 promoter activity. The activity of particular CXCR7 promoter constructs was assayed by dual luciferase showing an important role of NF-κB transcription factors. Activity was measured on the basis of firefly/Renillla luciferase activity and then the equimolar fold of difference was counted. Results were expressed relative to a value of 1.0 for cells transfected with pGL4.72 vector and empty pGL4.10. Averages of duplicates from three independent experiments are shown. Columns, mean; bars, SEM. Frag 1-6 P < 0.05, as compared with empty vector. *, P < 0.05, fragments compared with NF-κBmut fragment and Frag7 (Mann-Whitney test). C. NF-κB to CXCR7 promoter. ChIP assays of the indicated cell lines showed that binding sites 0.03, 0.3, and 1.0 kb are required for full CXCR7 promoter activity. NF-κB–based regulation of the promoter seems to be identical in both cell lines.

NF-κB Is Required for CXCR7 Promoter Activity

In the next step, RD and RD/PAX3-FKHR cells were transfected by luciferase reporter gene constructs driven by those different CXCR7 promoter fragments (Fig. 4B). We noticed that the PAX3-FKHR protein somehow diminished CXCR7 promoter activity as seen for Frag 1, Frag 2, and Frag 5. The highest promoter activity was observed for a 528 bp fragment (Frag 5) that contains two NF-κB (N1 and N2) and proximal HRE binding sites (Fig. 4B). We noticed that whereas removal of distal NF-κB (N2) and HRE binding sites in this construct resulted in a significant decrease of promoter activity, removal of the proximal NF-κB (N1) binding site completely abolished its activity. Similarly, we observed almost complete inhibition of promoter activity when the proximal NF-κB binding site was mutated in the longest CXCR7 promoter fragment (NF-κBmut; Fig. 4B). Subsequently, we did ChIP analysis using anti–NF-κB antibodies confirmed so that NF-κB binds to three proximal NF-κB binding sites flanked by N1, N2, and N3 primers (Fig. 4C).

Furthermore, analysis of the promoter activity of Frag 3 and Frag 4 suggested the presence of a putative negative regulatory site(s) located between NF-κB (N2) and the distal HRE binding site (Fig. 4A and B). Because this part of the promoter contains a YY1 binding site, we hypothesize that this negative regulator of transcription may affect CXCR7 expression. Our RQ-PCR analysis revealed that in normoxic conditions, the YY1 mRNA level is approximately four times higher in PAX3-FKHR–expressing cells (Fig. 5A, black columns). Consistent with this observation, when the YY1 binding site was mutated in the CXCR4 promoter, we observed upregulation of promoter activity in RD but not RD/PAX3-FKHR cells (Fig. 5B). This further supports our data shown in Fig. 2C that the NRF-1-PAX3-FKHR interaction prevents the access of YY1 to the CXCR4 promoter. In contrast, mutation of the YY1 binding site in the CXCR7 promoter produced higher promoter activity in RD/PAX3-FKHR cells and, at the same time, did not affect CXCR7 promoter activity in the RD cell line (Fig. 5C). This could be explained by RD/PAX3-FKHR expressing YY1 at much higher level and that YY1 is negative regulator of CXCR7 expression in these cells. Thus, our data supports the negative involvement of YY1 in CXCR4 and CXCR7 promoter activities in RD and RD/PAX3-FKHR cells, respectively.

FIGURE 5.
FIGURE 5.

YY1 transcription factor negatively regulates CXCR4 and CXCR7 expression. A. PAX3-FKHR translocation upregulates YY1 mRNA expression. RQ-PCR shows ∼5-fold upregulation of YY1 mRNA expression after transfection with PAX3-FKHR construct. B. and C. YY1 transcription factor is responsible for the downregulation of CXCR4 and CXCR7 expression. Both cell lines were transiently transfected with wild-type CXCR4 (B) and CXCR7 (C) promoter fragments and fragments containing YY1 mutations. After 24 h, luciferase activity was measured and promoter activity was calculated as in Figs. 2B and 4B. *, P < 0.05, paired Student's t test.

Interestingly, we noticed that YY1 expression at the mRNA level, which is higher in normoxic conditions in RD/PAX3-FKHR cells (Fig. 5A; Supplementary Fig. S4), becomes upregulated in hypoxic conditions in RD cells and downregulated in RD/PAX3-FKHR cells (Fig. 5A, white columns).

Expression of CXCR4 and CXCR7 Receptors Is Differently Regulated in RD and RD/PAX3-FKHR Cells in Response to Hypoxia

It has been reported that hypoxia upregulates CXCR4 expression in several cell types (29-31). To address this issue better for RMS cells and to determine whether CXCR7 expression is also modulated by oxygen level, we evaluated the expression of both receptors in RD and RD/PAX3-FKHR cell lines in normoxic and hypoxic conditions (Fig. 6A, right and left). In these experiments, RMS cells were exposed to hypoxia for 0 to 24 hours and expression of both receptors was evaluated by fluorescence-activated cell sorting and shown as mean fluorescence intensity (MFI). Figures 1A and 6A show that RD cells in normoxia express very low levels of CXCR4 and highly express CXCR7, but if transferred to hypoxic conditions, will highly upregulate CXCR4 and downregulate CXCR7 (Fig. 6A, right and left). In contrast, RD/PAX3-FKHR cells that in normoxia express high levels of CXCR4 and low levels of CXCR7 upregulate the expression of both receptors (Figs. 1 and 6A, right and left). Overall, these protein expression data correlated with changes at the mRNA level (data not shown). Because our data (Fig. 5B and C) indicated the involvement of YY1 in CXCR4 and CXCR7 promoter activity, we analyzed potential binding of YY1 to both promoters by using the ChIP assay (Fig. 6B). As expected, we noticed that in normoxic conditions in RD cells, YY1 binds to CXCR4 but not the CXCR7 promoter. In contrast, in RD/PAX3-FKHR cells, YY1 binds to CXCR7 but not the CXCR4 promoter. In hypoxic conditions, however, YY1 binds to the CXCR7 promoter in RD cells but does not interact with either promoter in RD/PAX3-FKHR cells (Fig. 6B). These differences in YY1 binding to the CXCR7 promoter in normoxic versus hypoxic conditions may explain the downregulation of CXCR7 expression on RD cells during hypoxia.

FIGURE 6.
FIGURE 6.

A. Effects of hypoxia on CXCR4 (left) and CXCR7 receptor (right) membrane expression in RD and RD/PAX3-FKHR cell lines. Hypoxia enhances the expression of CXCR4 in both RD and RD/PAX3-FKHR cells. At the same time, CXCR7 is upregulated during hypoxia in RD/PAX3-FKHR cells, but downregulated in RD cell. Results are the mean of three experiments. *, P < 0.05, versus cells cultured in normoxia (paired Student's t test). MFI ± SD. B. YY1 binding to CXCR4 and CXCR7 promoters is divergently regulated by hypoxia. RD and RD/PAX3-FKHR cells were subjected to normoxia and hypoxia for 16 h, and subsequent ChIP assays were done. PCR products were visualized on gels. C. CXCR4 expression in hypoxia is dependent on HIF-1a. Sequential deletions of CXCR4 promoter reveals the vast importance of the HRE binding site located 1.3 kb from the transcriptional start. When a promoter fragment devoid of this HRE was used, promoter activity was abolished. Fold of difference was calculated as in Fig. 2B. Columns, mean; bars, SEM. *, P < 0.05, activities of Frag3.1, Frag4, and HREmut as compared with full-length promoter fragments (Mann-Whitney test). D. CXCR7 expression in hypoxia is dependent on HIF-1a only in the RD/PAX3-FKHR cell line. Specific constructs lacking HRE binding sites were ligated with pGL4 vector and luciferase activity was measured. Depicted is the fold difference of activity relative to Renilla vector and control. Fold of difference was calculated exactly as in Fig. 2B. Columns, mean; bars, SEM. *, P < 0.05, activities of Frag6 and HREmut as compared with full-length promoter fragments (Mann-Whitney test).

Next, because NRF-1 and NF-κB are required in RMS cells for CXCR4 and CXCR7 activity, respectively, we perturbed the expression of both transcription factors by using NRF-1 shRNA or BAY 11-7082 (small molecular NF-κB inhibitor) in RD and RD/PAX3-FKHR cells (Tables 1 and 2). In addition, we did similar studies in the human PAX-3-FKHR+ ARMS cell line RH30. As expected, perturbation of NRF-1 expression by shRNA (∼80% of basic level) led to the inhibition of CXCR4 in RD/PAX3-FKHR cells that highly express CXCR4 (Fig. 1; Table 1). CXCR4 was also downregulated in PAX3-FKHR+ RH30 cells. In addition, as expected, perturbation of NRF-1 expression during hypoxia impaired the upregulation of CXCR4 in RMS cells (Table 1).

View this table:
Table 1.

Inhibition of NRF-1 on CXCR4 and CXCR7 expression in RD, RD/PAX3-FKHR, and RH30 rhabdomyosarcoma cells

View this table:
Table 2.

Inhibition of NF-κB on CXCR4 and CXCR7 expression in RD, RD/PAX3-FKHR, and RH30 rhabdomyosarcoma cells

In the next set of experiments, we found that inhibition of NF-κB activity downregulated CXCR7 but not CXCR4 expression both in normoxic and hypoxic conditions in RD and RD/PAX3-FKHR cells (Table 2). Of note, because PAX3-FKHR+ RH30 cells express CXCR7 at a very low level, no significant changes in CXCR7 expression in this cell line were observed.

Subsequently, to better assess the role of HRE binding sites in the expression of both receptors in hypoxic conditions, we did analysis of promoter activities in hypoxia using different CXCR4 and CXCR7 fragments (Fig. 6C and D) in which HRE fragments (indicated in Figs. 2A and 4A) were deleted or mutated. While analyzing CXCR4 promoter fragment activity, we noticed that upregulation of CXCR4 promoter activity was abolished in response to hypoxia in both RD and RD/PAX3-FKHR cells (Fig. 6C) when three proximal HRE binding sites were deleted (Fig. 2A, Frag 4). We observed a similar response when one distal HRE motif from these three putative binding sites was deleted (Fig. 2A, Frag 3.1) or mutated (Fig. 2A, HREmut) as shown in Fig. 6C. This shows that the HRE binding at the −1282 to −1286 site is crucial for CXCR4 upregulation under hypoxic conditions.

Similar experiments were done to address the role of putative HRE fragments (Fig. 4A) in CXCR7 expression (Fig. 6D). We noticed that deletion of both distal HRE binding sites did not affect promoter activity under hypoxia. However, deletion of the proximal HRE binding site (Fig. 4A, Frag 6) or its mutation (Fig. 4A, HREmut) prevented the upregulation of CXCR7 promoter activity under hypoxic conditions (Fig. 6D).

Next, the changes in CXCR4 and CXCR7 expression under hypoxic conditions were confirmed by functional chemotactic assays in response to SDF-1 and I-TAC (Fig. 7). It is known that whereas CXCR4 binds SDF-1 only, CXCR7 is activated by both SDF-1 and I-TAC (20). As predicted from promoter activity/expression studies during hypoxia (Fig. 6), RD cells that upregulate CXCR4 during hypoxia and downregulate CXCR7 (Fig. 6A, right and left) slightly enhance migration to SDF-1 and decrease migration to I-TAC (Fig. 7A). On the other hand, RD/PAX3-FKHR cells that upregulate expression of both receptors under hypoxia (Fig. 6A, right and left), enhance chemotaxis to both ligands (Fig. 7B). Finally, our chemotaxis results were confirmed in functional chemotactic assays in which we perturbed the expression of CXCR4 and CXCR7 in RMS cells by using shRNA (NRF-1) or BAY 11-7082 (NF-κB), respectively (Supplementary Fig. S3).

FIGURE 7.
FIGURE 7.

Effect of hypoxia on chemotactic response of RD (A) and RD/PAX3-FKHR (B) cells. Cells were incubated for 16 h in hypoxia in the presence of either SDF-1 (300 ng/mL) or I-TAC (100 ng/mL). Results are mean data of three experiments. *, P < 0.05, versus cells cultured in normoxia (paired Student's t test).

Discussion

RMS is the most common soft tissue sarcoma of adolescence and childhood, and clinical evidence suggests that ARMS is more aggressive and metastatic than ERMS (11, 12). Because of the highly metastatic ARMS phenotype, which is responsible for poor clinical prognosis, there is an urgent need to better identify the mechanisms that control the metastatic behavior of these cells and to develop effective antimetastatic treatment strategies to improve the survival of patients with RMS. We became interested in the role of two G protein–coupled seven transmembrane–span chemokine receptors, CXCR4 (17) and CXCR7 (21) in RMS metastasis.

In our previous work, we showed that CXCR4 receptor plays a crucial role in SDF-1–mediated metastasis of RMS cells to SDF-1–expressing organs such as bone marrow and lymph nodes (17). Accordingly, we reported that CXCR4 expression correlates with the more metastatic phenotype of RMS and, although SDF-1 did not affect the proliferation or survival of RMS cells, it induced locomotion and directional chemotaxis in several RMS cell lines (17).

With the recent identification of CXCR7, a new receptor for SDF-1 that also binds the I-TAC chemokine, we became interested in the role of the CXCR7-SDF-1 and CXCR7-I-TAC axes in RMS progression (21). We noticed that CXCR7, in contrast to CXCR4, is expressed at a high level on ERMS lines. Although signaling from activated CXCR7 was not associated with increased RMS proliferation or cell survival similarly to CXCR4, it plays an important role in adhesion and migration of RMS cells (21).

The studies described in this work were aimed at better understanding how the expression of both receptors is regulated at the promoter level in human RMS cells. Promoter analysis was done in ERMS line RD and ARMS-like PAX3-FKHR–transduced RD cells. We were also interested in addressing how hypoxia influences the expression of these genes in PAX3-FKHR–negative and PAX3-FKHR–positive RD cells. We identified minimal promoter regions for CXCR4 and CXCR7, obtained information on molecular regulation of both promoters, and provided evidence that expression of both promoters is differently regulated in RMS cells.

First, we defined a minimal CXCR4 promoter fragment and found that the NRF-1 binding site residing in the proximal promoter sequence plays a crucial role in CXCR4 receptor expression in human RMS cells. Based on the literature there were some transcription factors postulated as positive regulators of CXCR4 promoter transcriptional activity including cyclic AMP responsive element, NF-κB, and HGF (32-36). However, our work is congruent with a previous report indicating the NRF-1 binding site defines a minimal CXCR4 promoter fragment and that NRF-1 plays a crucial role in CXCR4 expression in human RMS cells (28, 37). In addition to promoter deletion and NRF-1 binding site mutagenesis studies, the requirement of NRF-1 for CXCR4 promoter activity was additionally supported in our present study after blocking its expression by using NRF-1 shRNA.

Next, we identified a minimal CXCR7 promoter fragment and provided evidence that the proximal NF-κB binding motif was required for its activity. The requirement of NF-κB for CXCR7 promoter activity was additionally supported in this study after exposing RMS cells to BAY 11-7082, which is a small molecular NF-κB inhibitor. Interestingly, the NF-κB binding site was also identified in the proximal region of the CXCR4 promoter and, as mentioned above, it was even postulated to play a role in CXCR4 expression in human breast cancer cells (32). However, our promoter deletion studies revealed that this transcription factor does not play a significant role in basic CXCR4 expression, at least in RMS cells.

We also found that expression of CXCR4 is positively modulated in RMS cells by PAX3-FKHR protein. It is known that this fusion protein is a much stronger transcription factor as compared with wild-type PAX3 (13). Our results support this and show that PAX3-FKHR enhances CXCR4 expression in a PAX3 binding site–dependent manner. Furthermore, some promoter activity was observed in RD/PAX3-FKHR cells but not in PAX3-expressing RD cells, even if the NRF-1 site was mutated. To support this latter observation, we did a ChIP analysis and showed that the PAX3-FKHR protein may in fact directly interact with NRF-1 protein. However, further mutagenesis studies are needed to identify which part of the NRF-1 molecule is involved in this protein-protein interaction.

We also noticed that PAX3-FKHR, while enhancing CXCR4 expression, somehow downregulated the expression of CXCR7 in RMS cells. This is in agreement with our previous studies showing that CXCR4 is highly expressed on PAX3/7-FKHR–expressing ARMS cells (17). This is in contrast with CXCR7 expression, which is higher on PAX3/7-FKHR–negative ERMS cell lines (21). A possible explanation of this differential regulation of both promoters by PAX3-FKHR could be explained by YY1 activity that, as reported, is a negative regulator of CXCR4 expression (38, 39).

However, the YY1 binding motif is present in both promoters, in case of CXCR4, in which the YY1 binding site is located between the proximal NRF-1 and PAX3 binding motifs, direct interaction between PAX3-FKHR-NRF-1 may prevent access of YY1 to its binding sequence in PAX3-FKHR+ cells. In contrast, analogical sequence in CXCR7 promoter is accessible for YY1 binding. This is a better explanation as to why CXCR4 is expressed at a higher level compared with CXCR7 in RD/PAX3-FKHR cells. To support this further, when the YY1 binding site was mutated in the CXCR4 promoter, we observed the upregulation of promoter activity in RD but not in RD/PAX3-FKHR cells. In contrast, the involvement of YY1 in regulating CXCR7 promoter activity is more complex. One of the factors affecting its role is the total level of YY1, which is higher in PAX3-FKHR–expressing cells. This explains why CXCR7 seems to be negatively affected by YY1 in RD/PAX3-FKHR cells that express YY1, as we showed, at a much higher level.

Hypoxia has an important effect on the expression of several genes that contain HRE motifs. It was reported that expression of CXCR4 is upregulated in several cell types in response to hypoxic conditions (29-31). Here, we provide evidence that this is true for CXCR4+ RMS cells as well. To support this, we showed that RD cells expressing very low levels of CXCR4 and even RD/PAX3-FKHR cells that already highly express this receptor, both upregulate its expression in response to hypoxia. Of note, we identified one of the HRE elements located at the −1282 to −1286 site in the CXCR4 promoter sequence as being crucial for hypoxia-induced upregulation of this receptor.

Similar studies on CXCR7 expression in RD cells revealed that it is downregulated during hypoxia. We envision that the downregulation of CXCR7 expression by hypoxia on ERMS cells gives SDF-1 an advantage to more robustly engage the CXCR4 receptor, which becomes upregulated during hypoxic conditions. Therefore, because CXCR4-SDF-1 signaling increases the motility of RMS cells (17), ERMS cells may become more metastatic in response to low oxygen levels. In contrast to RD cells, and as shown in this report, RD/PAX3-FKHR cells upregulate CXCR7 expression during hypoxia. Because these cells already express high levels of CXCR4 in normoxic conditions, additional upregulation of CXCR7 may play an important role in affecting some other prometastatic SDF-1–mediated properties of ARMS cells such as secretion of metalloproteinases or the tethering of migrating RMS cells in potential metastatic sites.

Of note, our data also supports the role of YY1 in regulating CXCR4 and CXCR7 expression during hypoxia. We noticed that YY1 expression, which is lower in normoxia in RD cells as compared with RD/PAX3-FKHR, becomes upregulated in hypoxia in RD cells and at the same time is downregulated in RD/PAX3-FKHR cells. Based on this, whereas hypoxia-induced upregulation of YY1 expression in RD cells correlates with the downregulation of CXCR7 levels, hypoxia-induced downregulation of YY1 expression in RD/PAX3-FKHR cells explains the increase in CXCR7 expression on these cells.

In conclusion, in this report for the first time, we analyzed the regulation of promoter activity at the molecular level of newly identified SDF-1–binding receptor CXCR7 and provide evidence on the pivotal role of a proximal NF-κB binding site in this process. We also provide novel data on the role of NRF-1 in CXCR4 expression in RMS cells, and the involvement of the PAX3-FKHR-NRF-1 complex in enhancing CXCR4 expression in more metastatic PAX3-FKHR+ ARMS cells. We also postulate that the interaction of PAX3-FKHR and NRF-1 proteins in the proximal part of the CXCR4 promoter in PAX3-FKHR+ ARMS cells prevents access of the negative regulator of transcription YY1 to its binding site. Furthermore, we report that both of these prometastatic receptors are differently regulated in human RMS cells during hypoxia. Overall changes in the receptor expression pattern during hypoxic conditions suggest that ERMS cells may become more metastatic as a result of enhanced CXCR4 expression that primarily governs SDF-1–dependent RMS motility.

Disclosure of Potential Conflicts of Interest

No potential conflicts of interest were disclosed.

Acknowledgments

Grant Support: NIH grants R01 CA106281-01 and R01 DK074720, the Henry M. and Stella M. Hoenig Endowment (M.Z. Ratajczak); NIH grants R01 CA64202 and R01 CA104896 (F.G. Barr); and NIH grant P20RR018733 from the National Center for Research Resources (M. Kucia).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Footnotes

  • Note: Supplementary data for this article are available at Molecular Cancer Research Online (http://mcr.aacrjournals.org/).

    • Received June 12, 2009.
    • Revision received October 28, 2009.
    • Accepted November 11, 2009.

References

  1. ↵
    1. Barr FG,
    2. Galili N,
    3. Holick J,
    4. Biegel JA,
    5. Rovera G,
    6. Emanuel BS
    . Rearrangement of the PAX3 paired box gene in the paediatric solid tumour alveolar rhabdomyosarcoma. Nat Genet 1993;3:113–7.
    OpenUrlCrossRefPubMed
    1. Davis RJ,
    2. Barr FG
    . Fusion genes resulting from alternative chromosomal translocations are overexpressed by gene-specific mechanisms in alveolar rhabdomyosarcoma. Proc Natl Acad Sci U S A 1997;94:8047–51.
    OpenUrlAbstract/FREE Full Text
    1. Davis RJ,
    2. D'Cruz CM,
    3. Lovell MA,
    4. Biegel JA,
    5. Barr FG
    . Fusion of PAX7 to FKHR by the variant t(1;13)(p36;q14) translocation in alveolar rhabdomyosarcoma. Cancer Res 1994;54:2869–72.
    OpenUrlAbstract/FREE Full Text
    1. Hazelton BJ,
    2. Houghton JA,
    3. Parham DM,
    4. et al
    . Characterization of cell lines derived from xenografts of childhood rhabdomyosarcoma. Cancer Res 1987;47:4501–7.
    OpenUrlAbstract/FREE Full Text
    1. Ruymann FB,
    2. Newton WA,
    3. Ragab AH,
    4. Donaldson MH,
    5. Foulkes M
    . Bone marrow metastases at diagnosis in children and adolescents with rhabdomyosarcoma. A report from the Intergroup Rhabdomyosarcoma Study. Cancer 1984;53:368–73.
    OpenUrlCrossRefPubMed
    1. Sandberg AA,
    2. Stone JF,
    3. Czarnecki L,
    4. Cohen JD
    . Hematologic masquerade of rhabdomyosarcoma. Am J Hematol 2001;68:51–7.
    OpenUrlCrossRefPubMed
    1. Sharp R,
    2. Recio JA,
    3. Jhappan C,
    4. et al
    . Synergism between INK4a/ARF inactivation and aberrant HGF/SF signaling in rhabdomyosarcomagenesis. Nat Med 2002;8:1276–80.
    OpenUrlCrossRefPubMed
    1. Collins MH,
    2. Zhao H,
    3. Womer RB,
    4. Barr FG
    . Proliferative and apoptotic differences between alveolar rhabdomyosarcoma subtypes: a comparative study of tumors containing PAX3-FKHR or PAX7-FKHR gene fusions. Med Pediatr Oncol 2001;37:83–9.
    OpenUrlCrossRefPubMed
    1. Bennicelli JL,
    2. Advani S,
    3. Schafer BW,
    4. Barr FG
    . PAX3 and PAX7 exhibit conserved cis-acting transcription repression domains and utilize a common gain of function mechanism in alveolar rhabdomyosarcoma. Oncogene 1999;18:4348–56.
    OpenUrlCrossRefPubMed
  10. ↵
    1. Kelly KM,
    2. Womer RB,
    3. Barr FG
    . PAX3-FKHR and PAX7-FKHR gene fusions in rhabdomyosarcoma. J Pediatr Hematol Oncol 1998;20:517–8.
    OpenUrlCrossRefPubMed
  11. ↵
    1. Anderson J,
    2. Ramsay A,
    3. Gould S,
    4. Pritchard-Jones K
    . PAX3-FKHR induces morphological change and enhances cellular proliferation and invasion in rhabdomyosarcoma. Am J Pathol 2001;159:1089–96.
    OpenUrlPubMed
  12. ↵
    1. Gordon T,
    2. McManus A,
    3. Anderson J,
    4. et al
    . Cytogenetic abnormalities in 42 rhabdomyosarcoma: a United Kingdom Cancer Cytogenetics Group Study. Med Pediatr Oncol 2001;36:259–67.
    OpenUrlCrossRefPubMed
  13. ↵
    1. Fredericks WJ,
    2. Galili N,
    3. Mukhopadhyay S,
    4. et al
    . The PAX3-FKHR fusion protein created by the t(2;13) translocation in alveolar rhabdomyosarcomas is a more potent transcriptional activator than PAX3. Mol Cell Biol 1995;15:1522–35.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Zlotnik A,
    2. Yoshie O
    . Chemokines: a new classification system and their role in immunity. Immunity 2000;12:121–7.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Horuk R
    . Chemokine receptors. Cytokine Growth Factor Rev 2001;12:313–35.
    OpenUrlCrossRefPubMed
  16. ↵
    1. Kucia M,
    2. Reca R,
    3. Miekus K,
    4. et al
    . Trafficking of normal stem cells and metastasis of cancer stem cells involve similar mechanisms: pivotal role of the SDF-1-CXCR4 axis. Stem Cells 2005;23:879–94.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Libura J,
    2. Drukala J,
    3. Majka M,
    4. et al
    . CXCR4-SDF-1 signaling is active in rhabdomyosarcoma cells and regulates locomotion, chemotaxis, and adhesion. Blood 2002;100:2597–606.
    OpenUrlAbstract/FREE Full Text
    1. Muller A,
    2. Homey B,
    3. Soto H,
    4. et al
    . Involvement of chemokine receptors in breast cancer metastasis. Nature 2001;410:50–6.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Hartmann TN,
    2. Burger JA,
    3. Glodek A,
    4. Fujii N,
    5. Burger M
    . CXCR4 chemokine receptor and integrin signaling co-operate in mediating adhesion and chemoresistance in small cell lung cancer (SCLC) cells. Oncogene 2005;24:4462–71.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Burns JM,
    2. Summers BC,
    3. Wang Y,
    4. et al
    . A novel chemokine receptor for SDF-1 and I-TAC involved in cell survival, cell adhesion, and tumor development. J Exp Med 2006;203:2201–13.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Grymula K,
    2. Tarnowski M,
    3. Maksym R,
    4. Kucia M,
    5. Ratajczak J,
    6. Ratajczak MZ
    . An in vitro and in vivo evidence that human rhabdomyosarcomas (RMS) express functional CXCR7 receptor: overlapping and distinct role of CXCR4-SDF-1 and CXCR7-SDF-1/ITAC axes in regulating metastatic behavior of RMS cells. Blood 2008 (ASH Annual Meeting Abstracts) 112:4732.
    OpenUrl
  22. ↵
    1. Jankowski K,
    2. Kucia M,
    3. Wysoczynski M,
    4. et al
    . Both hepatocyte growth factor (HGF) and stromal-derived factor-1 regulate the metastatic behavior of human rhabdomyosarcoma cells, but only HGF enhances their resistance to radiochemotherapy. Cancer Res 2003;63:7926–35.
    OpenUrlAbstract/FREE Full Text
    1. Hartmann TN,
    2. Grabovsky V,
    3. Pasvolsky R,
    4. et al
    . A crosstalk between intracellular CXCR7 and CXCR4 involved in rapid CXCL12-triggered integrin activation but not in chemokine-triggered motility of human T lymphocytes and CD34+ cells. J Leukoc Biol 2008;84:1130–40.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Wysoczynski M,
    2. Miekus K,
    3. Jankowski K,
    4. et al
    . Leukemia inhibitory factor: a newly identified metastatic factor in rhabdomyosarcomas. Cancer Res 2007;67:2131–40.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Tomescu O,
    2. Xia SJ,
    3. Strezlecki D,
    4. et al
    . Inducible short-term and stable long-term cell culture systems reveal that the PAX3-FKHR fusion oncoprotein regulates CXCR4, PAX3, and PAX7 expression. Lab Invest 2004;84:1060–70.
    OpenUrlCrossRefPubMed
  26. ↵
    1. Miao Z,
    2. Luker KE,
    3. Summers BC,
    4. et al
    . CXCR7 (RDC1) promotes breast and lung tumor growth in vivo and is expressed on tumor-associated vasculature. Proc Natl Acad Sci U S A 2007;104:15735–40.
    OpenUrlAbstract/FREE Full Text
  27. ↵
    1. Moriuchi M,
    2. Moriuchi H,
    3. Fauci AS
    . HTLV type I Tax activation of the CXCR4 promoter by association with nuclear respiratory factor 1. AIDS Res Hum Retroviruses 1999;15:821–7.
    OpenUrlCrossRefPubMed
  28. ↵
    1. Wegner SA,
    2. Ehrenberg PK,
    3. Chang G,
    4. Dayhoff DE,
    5. Sleeker AL,
    6. Michael NL
    . Genomic organization and functional characterization of the chemokine receptor CXCR4, a major entry co-receptor for human immunodeficiency virus type 1. J Biol Chem 1998;273:4754–60.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Schioppa T,
    2. Uranchimeg B,
    3. Saccani A,
    4. et al
    . Regulation of the chemokine receptor CXCR4 by hypoxia. J Exp Med 2003;198:1391–402.
    OpenUrlAbstract/FREE Full Text
    1. Staller P,
    2. Sulitkova J,
    3. Lisztwan J,
    4. Moch H,
    5. Oakeley EJ,
    6. Krek W
    . Chemokine receptor CXCR4 downregulated by von Hippel-Lindau tumour suppressor pVHL. Nature 2003;425:307–11.
    OpenUrlCrossRefPubMed
  31. ↵
    1. Wang X,
    2. Li C,
    3. Chen Y,
    4. et al
    . Hypoxia enhances CXCR4 expression favoring microglia migration via HIF-1α activation. Biochem Biophys Res Commun 2008;371:283–8.
    OpenUrlCrossRefPubMed
  32. ↵
    1. Helbig G,
    2. Christopherson KW 2nd,
    3. Bhat-Nakshatri P,
    4. et al
    . NF-κB promotes breast cancer cell migration and metastasis by inducing the expression of the chemokine receptor CXCR4. J Biol Chem 2003;278:21631–8.
    OpenUrlAbstract/FREE Full Text
    1. Cristillo AD,
    2. Bierer BE
    . Regulation of CXCR4 expression in human T lymphocytes by calcium and calcineurin. Mol Immunol 2003;40:539–53.
    OpenUrlCrossRefPubMed
    1. Matteucci E,
    2. Locati M,
    3. Desiderio MA
    . Hepatocyte growth factor enhances CXCR4 expression favoring breast cancer cell invasiveness. Exp Cell Res 2005;310:176–85.
    OpenUrlCrossRefPubMed
    1. De La Luz Sierra M,
    2. Gasperini P,
    3. McCormick PJ,
    4. Zhu J,
    5. Tosato G
    . Transcription factor Gfi-1 induced by G-CSF is a negative regulator of CXCR4 in myeloid cells. Blood 2007;110:2276–85.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Cristillo AD,
    2. Highbarger HC,
    3. Dewar RL,
    4. Dimitrov DS,
    5. Golding H,
    6. Bierer BE
    . Up-regulation of HIV coreceptor CXCR4 expression in human T lymphocytes is mediated in part by a cAMP-responsive element. FASEB J 2002;16:354–64.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Moriuchi M,
    2. Moriuchi H,
    3. Turner W,
    4. Fauci AS
    . Cloning and analysis of the promoter region of CXCR4, a coreceptor for HIV-1 entry. J Immunol 1997;159:4322–9.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Moriuchi M,
    2. Moriuchi H,
    3. Margolis DM,
    4. Fauci AS
    . USF/c-Myc enhances, while Yin-Yang 1 suppresses, the promoter activity of CXCR4, a coreceptor for HIV-1 entry. J Immunol 1999;162:5986–92.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Hasegawa A,
    2. Yasukawa M,
    3. Sakai I,
    4. Fujita S
    . Transcriptional down-regulation of CXC chemokine receptor 4 induced by impaired association of transcription regulator YY1 with c-Myc in human herpesvirus 6-infected cells. J Immunol 2001;166:1125–31.
    OpenUrlAbstract/FREE Full Text
View Abstract
Back to top
Molecular Cancer Research: 8 (1)
January 2010
Volume 8, Issue 1

Sign up for alerts

View this article with LENS

Open full page PDF
Article Alerts
Email Article
Citation Tools
Regulation of Expression of Stromal-Derived Factor-1 Receptors: CXCR4 and CXCR7 in Human Rhabdomyosarcomas
Maciej Tarnowski, Katarzyna Grymula, Ryan Reca, Kacper Jankowski, Radoslaw Maksym, Joanna Tarnowska, Grzegorz Przybylski, Frederic G. Barr, Magdalena Kucia and Mariusz Z. Ratajczak
Mol Cancer Res January 1 2010 (8) (1) 1-14; DOI: 10.1158/1541-7786.MCR-09-0259
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

Advertisement