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[Cancer Research 60, 6160-6170, November 1, 2000]
© 2000 American Association for Cancer Research

Tumor Biology

Induction of Cell Death by Basic Fibroblast Growth Factor in Ewing’s Sarcoma1

Lisa-Marie Sturla, Georgina Westwood, Peter J. Selby, Ian J. Lewis and Susan A. Burchill2

Candlelighter’s Children’s Cancer Research Laboratory [L-M. S., G. W., S. A. B.], Imperial Cancer Research Fund Cancer Medicine Research Unit [P. J. S.], and Department of Pediatric Oncology [I. J. L.], St. James’s University Hospital, Leeds LS9 7TF, United Kingdom


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
Ewing’s sarcoma is thought to arise after developmental arrest of primitive neural cells during embryogenesis. Because basic fibroblast growth factor (bFGF) has a critical role in the regulation of cell survival, proliferation, and differentiation during embryogenesis, we have tested the hypothesis that bFGF and FGF receptors may contribute to the development of Ewing’s sarcoma and may provide a mechanism for the modulation of their behavior. All four of the Ewing’s sarcoma cell lines examined expressed bFGF and FGF receptors, which were detected by immunofluorescence and Western blotting. bFGF-induced a significant dose-dependent decrease in Ewing’s sarcoma cell proliferation on plastic and reduced anchorage-independent growth in soft agar. Unexpectedly, this decrease in cell number reflected bFGF-induced apoptosis and necrosis, as demonstrated by electron microscopy, binding of annexin V, and staining with acridine orange. Induction of cell death was dependent on dosage of, and period of exposure to, bFGF. bFGF did not induce differentiation of Ewing’s sarcoma cells in either the presence or the absence of serum or nerve growth factor. Treatment of NuNu mice with bFGF decreased growth of the highly tumorigenic Ewing’s sarcoma cell lines. Histologically tumors grown in the NuNu mice treated with bFGF were less cellular than those in control mice, and showed an increased level of apoptotic nuclei. This is in contrast to the mitogenic effect bFGF has in most other cancer cells. In summary, bFGF decreases Ewing’s sarcoma growth in vitro and in vivo by the induction of cell death. This novel observation may provide a new therapeutic strategy for Ewing’s sarcomas.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
bFGF3 belongs to a family of heparin-binding polypeptide growth factors and was originally identified in extracts of pituitary and brain tissue (1) . It is ubiquitously expressed but is most abundant in the nervous system (2) , affecting a broad spectrum of developmentally regulated cellular responses involved in the control of growth and differentiation (3 , 4) . Levels of bFGF are high during neuronal morphogenesis (5) , in which it has been shown to promote survival and repair of neurons (2 , 6) . This suggests that bFGF has an important role in maintaining specific neuronal populations (3) . bFGF commonly increases cell proliferation (7, 8, 9) , and inappropriate expression of this growth factor and its receptors has been implicated in transformation and malignant progression (10, 11, 12) . The use of bFGF to treat malignancy would, therefore, appear counterintuitive.

bFGF signal transduction occurs through a family of high- and low-affinity FGF receptors, which are thought to account for its diverse effects. Four high-affinity receptors sharing the same basic structure have been described, each containing an intracellular split-kinase domain and an extracellular domain containing up to three immunoglobulin-like domains. Structural variants of the high-affinity receptors can be generated by alternative splicing (13) , resulting in modified ligand binding (14 , 15) and subcellular localization (16) . These are expressed in a cell- and tissue-specific manner, which may change during lineage development (10 , 17) .

Tumors of the Ewing’s sarcoma family, including the peripheral primitive neuroectodermal tumors (pPNETs), are small round-cell tumors arising in the bone or soft tissues in persons predominantly between the ages of 10 and 20 years. The histogenic origin of Ewing’s sarcoma has been a matter of some dispute, although recent evidence confirms a primitive pluripotent neural cell of origin (18) . The variety of bony and soft tissue locations for these tumors may be explained in part by the wide distribution of pluripotent stem cells throughout the parasympathetic autonomic nervous system. Despite some improvements in treatment and outcome, less than 20% of patients who present with metastatic disease are long-term survivors, demonstrating the need for new treatment strategies.

For many cancers, including the neurally derived childhood tumor neuroblastoma (19) , histological and biochemical features of differentiation are associated with a good prognosis. This has lead to the evaluation of differentiation therapies, the aim being to selectively engage the process of terminal differentiation leading to restoration of normal cellular homeostasis. In neuroblastoma, in vitro studies have shown that treatment with agents such as NGF (20 , 21) or retinoic acid (22, 23, 24) induces differentiation, and more recently, the clinical efficacy of retinoic acid analogues has been demonstrated (25) . However, Ewing’s sarcomas appear to have lost the ability to engage terminal differentiation (18 , 26) . We, therefore, formed the hypothesis that because these tumors are derived from a primitive neural crest progenitor, they might be too immature to undergo differentiation after treatment with commonly used differentiation-inducing agents. If this were true, treatment of Ewing’s sarcoma cells with growth factors or hormones that commit pluripotent cells toward a differentiated lineage might modulate their behavior and response to differentiation-inducing agents. Such use of growth-promoting agents in the treatment of this aggressive malignancy has not previously been considered.

bFGF has a critical role in the commitment of primitive neural cells toward a neuronal phenotype (27 , 28) . Although exposure to NGF is an important mediator of neuronal differentiation, it will induce differentiation of sympathoadrenal progenitors only when the cells have first been exposed to bFGF (28) . To determine whether bFGF drives Ewing’s sarcoma cells toward a neural phenotype, which might paradoxically be exploited therapeutically, the effects of bFGF on their growth, survival, and differentiation in the presence and absence of NGF have been examined for the first time.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell Lines
The well-characterized Ewing’s sarcoma family cell lines RD-ES, TC-32, SK-N-MC, A673, and TTC-466 were studied (29) . All of the cell lines were derived from soft tissue peripheral primitive neuroectodermal tumors, with the exception of RD-ES, which was derived from a bony Ewing’s sarcoma. The TC-32 and RD-ES cell lines were maintained in RPMI 1640 (Sigma, Poole, United Kingdom) supplemented with 10% FCS (Seralab, Sussex, United Kingdom). SK-N-MC and A673 cells were maintained in DMEM nutrient mixture HAM F12 (Sigma) and DMEM, respectively, supplemented with 10% FCS. The neuroblastoma cell line IMR-32, used as a positive control in proliferation and soft agar studies, was cultured in DMEM/RPMI 1640 (Sigma) plus 10% FCS. The breast carcinoma cell line MCF-7 was used as a positive control for bFGF and FGF receptor studies, and was maintained in DMEM supplemented with 10% FCS. With the exception of the TC-32 and RD-ES cells, which were a kind gift from Dr. J. A. Toretsky (National Cancer Institute, Bethesda, MD), all of the cell lines were purchased from the American Tissue Culture Collection (Rockville, MD).

bFGF
Lyophilized bFGF (25 µg; Sigma) was dissolved in 1 ml of sterile PBS (pH 7.4) containing 1% (w/v) fatty-acid-free BSA (Sigma). bFGF was aliquoted and stored at -20°C until required. Biological activity of bFGF was assayed using a PC12 neurite extension assay (results not shown). For in vivo studies, bFGF was dissolved in normal growth media and delivered daily by a single s.c. injection.

Viable Cell Number
Cells (2 x 105) were seeded in Primaria 6-well plates and incubated in normal media with serum for 24 h. Media were removed by aspiration and replaced with fresh media plus 10% FCS alone or supplemented with bFGF (10–80 ng/ml). Cells were incubated at 37°C in a 95% air-5% CO2 humidified atmosphere for 24–96 h. In some experiments, cells were exposed to bFGF (20 ng/ml) for 6, 12, 24, or 48 h; media were removed and replaced with fresh media plus 10% FCS, followed by incubation for an additional 24, 48, or 72 h. After incubation, cells were harvested using EDTA (0.05%) and trypsin (0.1%), and viable cell numbers were counted using the trypan blue exclusion assay and a Neubauer hemocytometer. Results are shown as mean ± SE (n = 12; Fig. 3aCitation ).



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  		Fig. 3. bFGF decreases viable Ewing’s sarcoma cell number in a dose- and time-dependent manner. Exposure of Ewing’s sarcoma cells to bFGF (20 ng/ml) decreased the viable cell number. In a, after treatment with bFGF (20 ng/ml) for 48 h, there were significantly fewer viable cells in the control population compared with the bFGF-treated cells at 24 (P < 0.01), 48 (P < 0.0001), and 72 h (P < 0.0001). Results are shown for viable TC-32 cell number at 0, 24, 48, and 72 h in control cells ({diamondsuit}) and in cells treated with bFGF (20 ng/ml) for 48 h ({blacksquare}). In b, exposure to bFGF for 6, 12, 24, or 48 h decreased the viable cell population to ~50% of that in the untreated cell population at 24 h. After returning cells to normal growth conditions for 24 h, there was no significant difference in viable cell number in cells treated with bFGF ({blacksquare}) at 6, 12, 24, or 48 h, which suggests that these cells are not proliferating. However the control ( ) untreated cultures continued to increase in number. Results are shown as mean ± SE (n = 6). Statistical analysis was made by ANOVA, with a Bonferroni t test. Ps are shown for viable cell number in control and bFGF-treated cultures.

 
BrdUrd Proliferation Assay
Cells (1.5 x 103 ) were seeded in Primaria 96-well plates and incubated in normal medium with serum for 24 h. Medium was removed, and wells were rinsed with a serum-free medium before treating with bFGF (2 pg–2 µg/ml) in normal medium supplemented with 10% FCS or in a serum-free defined medium containing sodium selenite (30 nM), progesterone (20 nM), putrescine (100 µM), transferrin (100 µg/ml), and insulin (10 µg/ml). Cells were incubated at 37°C in a 95% air-5% CO2 humidified atmosphere.

The Biotrak cell proliferation ELISA system, version 2 (Amersham) was used to measure proliferation, following manufacturer’s instructions. BrdUrd (13.3 µM) was added to culture medium 2 h prior to assay of cells at 24, 48, and 72 h. Absorbance (Abs) was determined at 450 nm (Titertek Multiskan plate reader). Relative BrdUrd incorporation is shown as (absorbance value x 1000) ± SE (n = 3). Each assay was repeated three times, with 7 replicates of each condition per assay.

Soft Agar Tumorgenicity Assay
Primaria Petri dishes (35-mm) were coated with 0.5 ml of endotoxin free agar (0.3% v/v; Life Technologies, Inc., Paisley, United Kingdom) to prevent cells adhering to the bottom of the dish. A single-cell suspension (5 x 104 cells/ml) of each cell line was prepared in serum-supplemented medium with bFGF (10 or 20 ng/ml), and agar was added to a final concentration of 0.3% (w/v) before plating 1 ml of cell suspension into the precoated Petri dishes. Agar was allowed to set for 10 min, and dishes were placed in a 10-cm2 Petri dish containing one 35-mm dish of sterile ddH2O to maintain humidity. Dishes were incubated at 37°C in a humidified atmosphere of 95% air-5% CO2, and colony formation was observed over 14 days. Colony number and size were scored in four randomly selected fields for each dish. Each condition was tested in triplicate, and each assay was repeated at least three times.

Ewing’s Sarcoma Xenografts in Nude Mice
Equal numbers of RD-ES or TC-32 cells (2.5 x 106 suspended in 200 µl of growth media) were delivered by a single s.c. injection in the right flank of female NuNu mice. After 8 days, mice were treated with either bFGF (100 or 200 ng in 0.1 ml of growth media/mouse/day) or with 0.1 ml of medium alone. Mice were examined twice weekly for tumor growth, and palpable tumor size was recorded. When tumors reached approximately 1.4 cm3 , mice were killed, tumors were excised, and sizes were accurately measured before they were mounted in OCT and frozen in liquid nitrogen-cooled isopentane. Cryosections (10-µm) of tumors were prepared and stained with H&E. Endogenous peroxidase was quenched in sections by treating with hydrogen peroxide (3% v/v in PBS; BDH) for 5 min at room temperature. Expression of the proliferation marker Ki 67 was examined using a sheep anti-Ki 67 antibody (working concentrate, 1:200 dilution of antibody. The Binding Site, Birmingham, United Kingdom). Staining for Ki 67 was visualized using peroxidase-antiperoxidase, sections counterstained with H&E and mounted in DePeX mounting medium (BDH). The sections were examined by light microscopy using a Zeiss Axioplan microscope (x40), and the number of positive cells were scored in 5 fields per xenograft. The proliferation index = number of Ki 67-positive nuclei ÷ number of nuclei scored ( proliferation index: 1, all cells proliferating; 0, no cells proliferating). The number of apoptotic nuclei was scored using the TUNEL assay (see "TUNEL Assay" below).

Characterization of Cell Death
Light and Electron Microscopy.
Cells (2 x 106) were seeded in 75-cm2 flasks and were cultured under normal growth conditions for 24 h. The media were removed and replaced with media alone or media containing bFGF (20 ng/ml) for up to 4 days. Cells were harvested by gentle scraping with a rubber scraper and were centrifuged at 900 x g for 4 min to form a soft pellet. Cells were then fixed in 2.5% gluteraldehyde in Sorenson’s buffer [0.16 M disodium hydrogen Pi, 0.04 M sodium dihydrogen phosphate (pH7.4)] for 1 h before four 20-min washes in Sorenson’s wash buffer [80 mM disodium hydrogen Pi, 20 mM sodium dihydrogen phosphate (pH7.4)]. Sections (5-µm) were examined by light and electron microscopy. The number of apoptotic, mitotic, and necrotic cells per 1000 were scored (x1500). Only cells in which a nucleus could be seen were scored. Apoptotic bodies were not counted. Cells demonstrating increased overall and nuclear size, with breakdown of the nuclear and plasma membranes and loss of organelles, were scored as necrotic.

Annexin V Binding.
Cells were grown in 25-cm2 Primaria flasks and treated with bFGF (5–80 ng/ml) for up to 72 h. Cells were harvested by trypsinization and were resuspended in ice-cold RPMI-HEPES (10 mM; Sigma) at a density of 5 x 105 cells/ml. Time of exposure to EDTA (0.1% in PBS) and trypsin (1x in PBS) and pipetting were kept to a minimum to avoid cell damage. Cells were labeled with annexin V (1:200; Alexis) and PI (1 µg/ml; Sigma) for 30 min before preparation of cytospins (500 g for 5 min; Shandon Cytospin 3) or FACS (Becton Dickinson FACScan). Cytospins were viewed and photographed immediately by fluorescence microscopy using an Axioplan Zeiss microscope.

Acridine Orange Staining.
Cells were seeded on 12-well slides (5 x 10-3 cells/well) and, after 24 h, were treated with bFGF (20 ng/ml) for 0–4 days. Slides were fixed at 12-h intervals in methanol/acetone (1:1) and were stained with acridine orange (30 µg/ml; Sigma) in phosphate buffer [5 mM Na2HPO4 (pH 7.4)] for 10 s. Cells were washed three times for 10 min each in phosphate buffer and were visualized by fluorescence microscopy using an Axioplan Zeiss microscope.

TUNEL Assay.
Apoptosis in xenografts from control and bFGF-treated mice was determined using the TUNEL assay (30) to detect DNA fragmentation (ApoTag; Intergen, Oxford, United Kingdom). Briefly, cryostat sections (10 µm) were fixed in paraformaldehyde [1% w/v in PBS (pH 7.4)] for 10 min at room temperature, followed by a post-fix in precooled ethanol:acetic acid (2:1) for 5 min at -20°C. Endogenous peroxidase was quenched by treating sections with hydrogen peroxide (3% v/v in PBS) for 5 min at room temperature. DNA strand-breaks were detected by enzymatically labeling the free 3'-OH termini with modified nucleotides and staining with peroxidase-antiperoxidase according to the manufacturer’s instructions (ApoTag; Intergen). Sections were counterstained with crystal violet-free methyl green [0.5% w/v in 0.1 M sodium acetate (pH 4)], dehydrated in xylene (100%), and mounted in DePeX mounting medium. The sections were examined by light microscopy using a Zeiss Axioplan microscope (x40) and the number of apoptotic nuclei were scored in 5 fields per xenograft. The apoptotic index = number of apoptotic nuclei ÷ number of nuclei scored (apoptotic index: 1, all nuclei apoptotic; 0, no nuclei apoptotic.


   Western Blotting
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
Cells were grown in media alone and in media supplemented with bFGF (20 ng/ml) for neurofilament studies. Cells were harvested at intervals between 8 h and 4 days by trypsinization, and pellets were washed twice with PBS. Protein extracts were prepared in lysis buffer [1 M NaCl; 10 mM Tris-HCl (pH7.6); 1 mM EDTA; 1 µg/ml aprotinin, and 100 µg/ml phenylmethylsulfonylfluoride], and the protein content of cell extracts was estimated using the Bio-Rad DC protein assay (Bio-Rad Laboratories). Each protein sample (20 µg) was size-fractionated by SDS-polyacrylamide (10%) gel electrophoresis. The accuracy of protein estimation and loading was confirmed by staining the separated proteins using the Lowry silver stain (Pharmacia Biotech).

Proteins were transferred onto nitrocellulose membrane (Hybond-C; Amersham) in transfer buffer (25 mM Tris, 192 mM glycine, and 20% methanol) using a mini-transblot system (Bio-Rad Laboratories) overnight at 4°C. Membranes were blocked with a 5% nonfat milk solution in 1x TTBS [0.05% Tween 20, 20 mM Tris-HCl (pH7.5), and 500 mM NaCl] for 2 h. Membranes were incubated in rabbit polyclonal anti-pan-neurofilament and FGF receptor-1 antibodies [1:500 (Affiniti) and 1:20 (Santa Cruz), respectively) for 1 h in a 2% nonfat milk solution in 1x TTBS. The filter was then washed twice for 5 min in 1x TTBS and was incubated for 1 h in streptavidin-conjugated goat antirabbit antibody (1:2000; Sigma). Membranes were washed (three time for 5 min each) in 1x TTBS, and secondary antibody was detected by enhanced chemiluminescence (Hyperfilm ECL; Amersham).


   Immunofluorescence
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
Cells were cultured on 12-well glass slides at a density of 5 x 10-3 cells/well under normal culture conditions. After a 48-h incubation period, medium was removed, and cells were fixed in methanol/acetone (1:1) twice for 2 min each before they were air-dried. Cells were incubated with rabbit polyclonal anti-pan-neurofilament (1:500; Affiniti), anti-bFGF (20 µg/ml; Santa Cruz), or anti-FGF receptor-1 (200 µg/ml, Santa Cruz) antibodies for 1 h, followed by two washes with PBS and refixing in methanol/acetone twice for 2 min each. Cells were again air-dried before a 30-min incubation with FITC-conjugated goat antirabbit antisera (1:300; Sigma). Two washes in PBS and two washes in PBS-0.25% Tween 20 were followed by a final rinse in ddH2O and air-drying. Cells were mounted in DABCO-glycerol and viewed by fluorescence microscopy. Specificity of immunofluorescence was confirmed by the absence of staining in a primary antibody negative control.


   RT-PCR
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 
Primers designed to amplify the first immunoglobulin-like loop (I) of the extracellular domain or the transmembrane/kinase domains (K) of each of the four high-affinity FGF receptors were used (Table 1)Citation . PCR conditions were optimized using total RNA isolated from the MCF-7 breast carcinoma cell line. RNA (2 µg) was reverse transcribed at 37°C for 1 h using 15 units of murine Moloney leukemia virus reverse transcriptase (Pharmacia Biotech) in 1x PCR buffer, [10 mM Tris-HCl (pH 8.3) and 50 mM KCl (Perkin-Elmer, Warrington, United Kingdom)], 1 mM dNTP (Pharmacia Biotech), 10 mM MgCl2, 1.2 µg of random primers (Life Technologies, Inc.), and 28 units of RNA guard (Pharmacia Biotech). cDNA was divided to amplify all of the target FGF receptors. cDNA (10 µl) was amplified using 2.5 units of Amplitaq gold (Perkin-Elmer) and primers at a concentration of 40 pmol per reaction in 1x PCR buffer (as above), 0.2 mM dNTP, and 2 mM MgCl2. Activation of Amplitaq gold with one cycle of 95°C for 10 min was followed by amplification for 35 cycles of 95°C for 30 s, annealing at 55–68°C (primer dependent; Table 1Citation ) for 30 s, extension at 72°C for 45 s, and a final cycle of 72°C for 7 min. A MCF-7 positive control, and reverse transcriptase and water negative controls were included for each set of PCR primers. PCR products were separated in a 1.5% agarose gel and {phi}X174 RF DNA/HaeIII DNA fragments (Life Technologies, Inc.) were used as markers for product sizing. The identity of PCR products for all of the four receptors was confirmed by sequence analysis using an ABI 377 automated sequencer (ABI PRISM Big dye terminator kit; Perkin-Elmer).


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  		Table 1 Primers for RT-PCR of FGF receptors 1 to 4, first immunoglobulin-like loop (I) and transmembrane/tyrosine kinase domain (K)

 

   Statistical Analysis
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
RESULTS
 DISCUSSION
 REFERENCES
 
Statistical analysis was performed by one-way ANOVA with a Bonferroni, Dunnett, or paired Student t test. A P of <0.05 was considered significant.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
DISCUSSION
 REFERENCES
 
bFGF and FGF Receptors Are Ubiquitously Expressed by Ewing’s Sarcoma Cell Lines.
All of the Ewing’s sarcoma cell lines examined expressed bFGF as demonstrated by immunofluorescence (Fig. 1a)Citation . Immunofluorescence for bFGF showed diffuse staining throughout the cell, although expression was stronger in the cytoplasm than in the nucleus. Expression of FGF receptors was also evident in all of the Ewing’s sarcoma cell lines studied. Immunofluorescence using an FGF receptor-1 antibody showed diffuse staining throughout the cell (Fig. 1a)Citation . The same antibody used for Western blotting confirmed the presence of FGF receptor-1, detecting a protein of the expected Mr 145,000 size in all of the Ewing’s sarcoma cell lines and the breast carcinoma positive control (Fig. 1b)Citation . However, additional proteins were also detected with this antibody, possibly pertaining to the receptor at various stages of glycosylation (31) or cross-reactivity with related proteins.



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  		Fig. 1. Expression of bFGF and FGF receptors in Ewing’s sarcoma cell lines. All of the cell lines examined expressed bFGF and FGF receptors, demonstrated by immunofluorescence (a), Western blotting (b), and RT-PCR(c). a, immunofluorescence using an anti-pan bFGF or anti-FGF receptor-1 antibody. Both of the antibodies showed diffuse-staining throughout the cell, although bFGF showed some specific localization to the cell nucleus. Expression of bFGF and FGF receptors is shown in the TC-32 cell line. b, Western blotting with the same FGF receptor-1 antibody, demonstrating the presence of a Mr 145,000 band (145KDa)—consistent with expression of FGF receptor-1—in the breast carcinoma positive control MCF-7 (Lane 1) and the Ewing’s sarcoma cell lines, TC-32 (Lane 2), RD-ES (Lane 3), A673 (Lane 4), SK-N-MC (Lane 5), and TTC-466 (Lane 6). The correct-size band was not detected in the COS-7 negative control (Lane C). Additional proteins were also detected with this antibody, which may reflect expression of the receptor at different stages of glycosylation or cross-reactivity with related proteins. c, RT-PCR for the extracellular (I) or transmembrane kinase (K) domain of each of the four high-affinity FGF receptors. The Ewing’s sarcoma cell lines expressed all four of the high-affinity FGF receptors; the MCF-7 breast carcinoma cell line expressed full-length FGF receptors 1, 2, and 4, but FGF receptor 3 was modified in the transmembrane kinase domain. Two novel forms of FGF receptor-3, modified in the transmembrane kinase domain, were identified in the Ewing’s sarcoma cell lines by RT-PCR. Results for the MCF-7 breast carcinoma cell line (positive control, Lane a) and the Ewing’s sarcoma cell line TC-32 (Lane b) are shown. m, molecular weight markers.

 
In the absence of specific antibodies suitable for Western blotting or immunohistochemistry, RT-PCR using primers to the extracellular (I) or transmembrane kinase (K) domains of the FGF receptors was used to demonstrate the presence of all four of the high-affinity receptors in the Ewing’s sarcoma cell lines (Fig. 1c)Citation . In addition the Ewing’s sarcoma cell line, SK-N-MC and A673 expressed an apparently truncated form of FGF receptor-2, missing the first immunoglobulin-like loop of the extracellular domain. Two variant forms of FGF receptor-3 were also detected in the Ewing’s sarcoma cells and the MCF-7 breast carcinoma positive control; sequence analysis showed these variant forms to be lacking the second half of the third immunoglobulin-like loop of the extracellular domain and the transmembrane domain.

bFGF Decreases Ewing’s Sarcoma Cell Numbers in Vitro.
Under normal serum-supplemented growth conditions, all three of the Ewing’s sarcoma cell lines studied—TC-32, RD-ES, and SK-N-MC—showed a significant reduction in proliferation when treated with bFGF (2–20 ng/ml) for 48 h (ANOVA, F7160 = 22.12, 29.54, and 8.59 respectively; P < 0.0001 in all of the cases), as demonstrated by BrdUrd incorporation (Fig. 2)Citation . The RD-ES cell line demonstrated the greatest reduction in proliferation, with bFGF (2 ng/ml)-treated cultures exhibiting an 80% reduction in proliferation as compared with an untreated control (P < 0.0001, Dunnett’s t test). In the TC-32 and SK-N-MC cell lines, the reduction was 55 and 44%, respectively (P < 0.0001, Dunnett’s t test). A decrease in proliferation was seen after exposure of TC-32 and RD-ES cells to bFGF (5–80 ng/ml) in both serum-free defined medium and serum-supplemented medium (ANOVA, F5120 = 39.94 and 65.59, respectively; P < 0.0001). For example, RD-ES cells exhibited a 94% reduction in proliferation when cultured in serum-free defined medium supplemented with bFGF (5 ng/ml) for 48 h, not significantly different from the 89% reduction observed in serum-supplemented medium (P = 1.00, Bonferroni’s t test). At concentrations of >=200 ng/ml, bFGF had no effect on BrdUrd incorporation.



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  		Fig. 2. bFGF decreased the incorporation of BrdUrd. bFGF (2 pg–2 µg/ml) decreased the proliferation of Ewing’s sarcoma cell lines ({diamondsuit}, TC-32; {blacksquare}, RD-ES; x, SK-N-MC) under normal growth conditions. Proliferation was measured using an ELISA assay to detect the incorporation of BrdUrd, which is given as absorbance (Abs) units x 10-3. a, the effect of bFGF (2pg–2 µg) on BrdUrd incorporation. b, the effect of bFGF (2pg–2 µg) on BrdUrd incorporation as a percentage of control cultures. c, bFGF (5–80 ng/ml) decreased BrdUrd incorporation in serum-free defined media ({blacksquare}) and under normal growth conditions in the presence of serum ({diamondsuit}) to the same extent. Results are shown as mean ± SE (n = 3). Statistical analysis was made using ANOVA analysis and a Dunnett t test.

 
This decrease in BrdUrd incorporation correlated with a decrease in viable Ewing’s sarcoma cell number (Fig. 3aCitation ; P < 0.0001, Bonferroni’s t test). Exposure of TC-32 cells to bFGF (20 ng/ml) for 6 and 12 h led to a 54% decrease in viable cell number after a 24-h recovery period (Fig. 3bCitation and Table 2Citation ; P < 0.05, Bonferroni’s t test). The percentage of viable cells in the bFGF-treated culture decreased to 44 and 27%, respectively, after 24-h (P < 0.001, Bonferroni’s t test) and 48-h (P < 0.0001, Bonferroni’s t test) exposure (Fig. 3bCitation and Table 2Citation ). However the total viable cell number after treatment with bFGF for 6, 12, 24, or 48 h was not significantly different (Fig. 3b)Citation . This population of apparently bFGF-resistant cells had a reduced rate of growth, as demonstrated by the slope of the growth curves (Fig. 3a)Citation . Similar results were found in the RD-ES cell line (Table 2)Citation .


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  		Table 2 Effect of exposure to bFGF on cell number at 0, 24, 48 and 72 h after treatment

Effect of bFGF on TC-32 and RD-ES cell number. Results are shown as the percentage of viable cells in a cell population at 0, 24, 48, and 72 h after exposure to bFGF (20 ng/ml) for 6, 12, 24, or 48 h. Column of shadowed data shows the % of viable cells in TC-32 cultures 24 h after treatment with bFGF for 6 h, 12 h, 24 h, or 48 h; these data are also presented in the form of a histogram in Fig 3bCitation .

 
bFGF Inhibits Anchorage-independent Growth of Ewing’s Sarcoma Cells.
Both of the TC-32 and RD-ES cells formed colonies in 0.3% soft agar. The mean colony-forming efficiency of RD-ES cells was 4.8% and of TC-32 cells was 2.4%. Supplementing the growth medium with 10 ng/ml bFGF reduced the colony number by 68% in the TC-32 cells and by 87% in the RD-ES cells (Table 3Citation ; P < 0.0001, Dunnett’s t test). When the concentration of bFGF was doubled to 20 ng/ml, the reduction in colony formation was not significantly different from that observed with 10 ng/ml bFGF (P = 1.00, Dunnett’s t test). bFGF (10 and 20 ng/ml) also significantly reduced colony size (P < 0.0001, Dunnett’s t test). In marked contrast, treatment of the neuroblastoma cell line IMR-32 with bFGF increased the number and size of colonies formed in soft agar (P < 0.0001, Dunnett’s t test; Table 3Citation ), consistent with previous reports (11 , 12) .


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  		Table 3 Effect of bFGF, NGF, and bFGF plus NGF on anchorage-independent growth

Effect of bFGF on growth of TC-32 and RD-ES cells in soft agar. Cells were incubated in soft agar for 14 days. Four randomly selected fields per plate were visualized and photographed using a Zeiss light microscope, under dark field conditions and a magnification of x5. Prints (13 x 9 cm) were made, and colonies were counted and sized from these. IMR-32 cells were included as a positive control. Results are shown as mean colony number or size ± SE (n = 9). Where results are significantly different from the control by Dunnett’s t test, Ps are given.

 
bFGF Decreases Ewing’s Sarcoma Xenograft Growth in NuNu Mice.
s.c. injection of NuNu mice with RD-ES or TC-32 cells resulted in rapid tumor growth in all of the injected mice; <=30 days after s.c. injection with 2.5 x 106 tumor cells, 100% of the mice had grown tumors of 1.4 cm3 (Fig. 4)Citation and were killed according to approved protocols. However, the rate of tumor growth (assessed by the maximum area of palpable tumor) was significantly reduced in mice treated daily with bFGF (100 or 200 ng/mouse/day) 8 days after inoculation with tumor cells, compared with those injected with vehicle alone (P < 0.001, Bonferroni’s t test; Fig. 4Citation ).



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  		Fig. 4. bFGF reduces the rate of Ewing’s sarcoma xenograft growth in NuNu mice by the induction of apoptosis. Both RD-ES and TC-32 cell lines are highly tumorigenic, producing tumors of 1.4 cm3 in 100% of inoculated NuNu mice within 30 days. Treatment of mice (n = 10) with bFGF (100 ng) significantly decreased the rate of both RD-ES [P < 0.01 (a)] and TC-32 [P < 0.001 (b)] tumor growth compared with control-vehicle-only injected mice (n = 10). The decrease in RD-ES tumor growth was dose-dependent; treatment of mice (n = 10) with 200 ng of bFGF/mouse/day had a greater inhibitory effect than treatment with 100 ng/mouse/day. Statistical analysis was made by ANOVA, with a Bonferroni t test. a, {circ}, control; •, treated with bFGF (100 ng/kg/day); {blacktriangleup}, treated with bFGF (200 ng/kg/day); b, {square}, control; {blacksquare}, treated with bFGF (100 ng/kg/day).

 
Decreased Ewing’s Sarcoma Cell Number and Xenograft Growth Is Mediated by bFGF-induced Apoptosis and Necrosis.
bFGF (2–80 ng/ml) induced cell death in the three Ewing’s sarcoma cell lines that we studied in a time- and dose-dependent manner, as demonstrated by light- and electron microscopy, labeling with annexin V and PI, and staining of nucleic acids with acridine orange.

By electron microscopy and labeling with annexin V and PI, an increase in both necrotic and apoptotic cell number was seen in all of the Ewing’s sarcoma cells after exposure to bFGF (Fig. 5Citation and 6)Citation . Loss of membrane asymmetry, an early apoptotic marker, was detected by staining with annexin V after treatment with bFGF (2–80 ng/ml) for 24 h (Fig. 5a)Citation . FACS analysis for annexin V- and PI-labeled cells identified three cell populations: one labeling with annexin V alone; one with only PI; and one with both annexin V and PI (Fig. 5b)Citation . In exponentially growing control TC-32 and RD-ES cells, >90% of the cell population did not stain with PI or annexin V (Fig. 5b)Citation . However, 48 h after treatment of TC-32 and RD-ES cells with bFGF, 42 and 57% of the respective cell populations were either alive but apoptotic (stained with annexin V alone) or dead (stained with annexin V plus PI or with PI alone). After 72 h, 83% of the RD-ES and 66% of the TC-32 cell populations were dead (stained with annexin V and PI or with PI alone; Fig. 5bCitation ). At 72 h, there were no viable apoptotic cells.



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  		Fig. 5. Induction of apoptosis and necrosis in RD-ES cells treated with bFGF. In a, RD-ES cells treated with bFGF (20 ng/ml) for 36 and 48 h showed a decrease in viable cell number as seen under light microscopy. Loss of membrane asymmetry (an early apoptotic marker), detected by staining with annexin V, is shown after 36- and 48-h exposure to bFGF. At 36 and 48 h, bFGF-treated and control, untreated cultures showed a redistribution of staining with acridine orange. In b, under normal growth conditions, more than 90% of TC-32 and RD-ES cultures were viable (red, lower left quadrant) and did not stain with annexin V or PI. After treatment with bFGF (20 ng/ml), the number of viable apoptotic (blue, lower right quadrant) and dead cells (pink, upper right quadrant and green, upper left quadrant) was significantly increased. After 72 h exposure to bFGF, the majority of TC-32 and RD-ES cells were dead (pink or green upper quadrants; P < 0.0001).

 


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  		Fig. 6. Effect of bFGF on ultrastructure of RD-ES cells. Treatment of Ewing’s sarcoma cells with bFGF (20 ng/ml)-induced cell death. The ultrastructure of cells treated with bFGF showed evidence of both necrosis and apoptosis (a). The apoptotic cell number increased with increased exposure time to bFGF (b). In cultures exposed to bFGF for 4 days, it was not possible to score the apoptotic and necrotic cell number, the culture consisting largely of cell debris (results not shown). Counts are shown for approximately 1000 cells.

 
Less than 1% of the total cell population exhibited signs of cell death identified by electron microscopy in the Ewing’s sarcoma cells under normal growth conditions (Fig. 6)Citation . After exposure to bFGF (20 ng/ml) the necrotic and apoptotic cell population increased with time (Fig. 6)Citation . After a 4-day exposure to bFGF, there were very few live cells. The majority of cells were vacuolated and showed signs of apoptosis or necrosis. Cells that were scored as apoptotic showed typical margination of the chromatin and chromatin condensation. Cells had decreased in size and had become more electron-dense. The nuclear and plasma membranes remained intact, with the subsequent formation of apoptotic bodies. However, a population of cells with an intact nuclear membrane and chromatin margination, but with plasma membrane permeability, increased cytoplasmic volume and a loss of cellular organelles was also observed. These cells were scored as necrotic; consequently, the scoring of necrotic cells after electron microscopy is artificially high compared with the number of apoptotic cells.

Cryostat sections of RD-ES and TC-32 xenografts were highly cellular (Fig. 7aCitation , Control) and showed no evidence of DNA fragmentation (Fig. 7bCitation , Control). However, apoptotic nuclei were detected in the xenografts from mice treated with bFGF (Fig. 7bCitation , +bFGF), which suggests that the decreased rate of xenograft growth in bFGF-treated mice is associated with increased apoptosis. The apoptotic index in RD-ES and TC-32 xenografts was 0.005 ± 0.005 and 0.007 ± 0.001, respectively. After treatment of mice with 100 ng of bFGF/mouse/day, the apoptotic index of RD-ES xenografts significantly increased to 0.19 ± 0.03 (P < 0.0001, Student’s t test; 38-fold increase); in mice treated with 200 ng of bFGF, the apoptotic index increased to 0.28 ± 0.02 (P < 0.0001, Student’s t test; 56-fold increase). In TC-32 xenografts, bFGF (100 ng/mouse/day) increased the apoptotic index 20-fold (0.14 ± 0.04; P < 0.004, Student’s t test). Labeling with Ki 67 was reduced in the tumors of mice treated with bFGF (Fig. 7cCitation , +bFGF), compared with control-vehicle-only-treated mice. The proliferation index in bFGF-treated mice inoculated with RD-ES cells was 0.015 ± 0.005 compared with 0.29 ± 0.04 (P < 0.0001, Student’s t test) in control-vehicle-only-treated mice. Similar results were found in TC-32 xenografts (control, 0.23 ± 0.004; bFGF-treated, 0.012 ± 0.004; P < 0.0001, Student’s t test).



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  		Fig. 7. The number of apoptotic nuclei increases in xenografts from NuNu mice treated with bFGF. RD-ES and TC-32 xenografts from control and bFGF-treated NuNu mice were mounted in OCT and frozen through liquid nitrogen-cooled isopentane. Cryostat sections (10 µm) were fixed and stained with H&E, and viewed by light microscopy (a), stained for apoptotic nuclei using the TUNEL assay as described in the "Material and Methods" (b) or the Ki 67 proliferation antigen (c). Sections stained for Ki 67 were counterstained with H&E; the TUNEL-stained sections were counterstained with methyl green. Xenografts in control mice were very cellular (a); showed no, or very little evidence of, apoptosis (b); and had a high rate of proliferation (c). In contrast, bFGF-treated mice grew tumors that were reduced in size, less cellular (a), had fewer Ki 67-positive cells (c), and significantly more apoptotic nuclei than in the control group (b).

 
NGF Does Not Affect Ewing’s Sarcoma Cell Proliferation or Differentiation.
NGF (20 and 40 ng/ml) did not induce morphological differentiation in any of the Ewing’s sarcoma cell lines examined (Fig. 8a)Citation , unlike the neuroblastoma cell line IMR-32, in which treatment with NGF (20 ng/ml) increased neurofilament expression and induced neurite extension (results not shown) characteristic of neural differentiation. Substrate-dependent (Fig. 8b)Citation and -independent (Table 3)Citation proliferation of the Ewing’s sarcoma cell lines was unaffected by NGF (20 and 40 ng/ml) in the presence or absence of bFGF (20 ng/ml; P = 1.0 in all of the cases, Bonferroni’s t test).



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  		Fig. 8. NGF has no effect on Ewing’s sarcoma cell morphology (a) or on anchorage-dependent proliferation (b), in the presence or absence of bFGF. The morphology and proliferation of RD-ES cells after exposure to NGF (20 ng/ml) was unchanged compared with control, untreated cultures. Treatment with bFGF (20 ng/ml) for 24–72 h caused cells to round (a) and was associated with a decrease in proliferation, as measured by incorporation of BrdUrd (b). NGF (20–40 ng/ml) had no effect on the morphology or decrease in proliferation that was induced after exposure to bFGF (20 ng/ml). Results are shown as mean ± SE (n = 3). Statistical analysis was made by ANOVA with a Bonferroni t test correction. {blacksquare}, TC-32; , RD-ES; , SK-N-MC.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
REFERENCES
 
bFGF decreases cell number by inducing cell death in Ewing’s sarcoma. As far as we are aware, this is the first report of bFGF-induced cell death in neurally derived tumor cells. Previous studies in rat mesencephalic (32) and chick primitive neural (33) structures have shown bFGF to induce apoptosis, which supports the hypothesis that bFGF has an initial sorting role in early development. The induction of apoptosis by bFGF in Ewing’s sarcomas is, therefore, consistent with the hypothesis that these tumors arise in a primitive neural stem cell. The magnitude of bFGF-induced cell death was different in the cell lines studied, probably reflecting the heterogeneity of Ewing’s sarcomas. This suggests that Ewing’s sarcomas can arise from primitive cells at various stages, and that expression of different growth factors and hormones by Ewing’s sarcomas may modulate the effect of bFGF either directly (34) or by modifying the expression of FGF receptors (35) . Ewing’s sarcoma-derived cell lines demonstrated a biphasic response to bFGF, with concentrations of bFGF >200 ng/ml failing to significantly affect cell proliferation in vitro. This may reflect the activation of additional signaling pathways within the cell, counteracting the bFGF-induced cell death pathway, either directly through the FGF receptors or indirectly by interaction with other receptors. FGF receptor signaling is initiated after FGF-induced dimerization of the high-affinity FGF receptors, which is regulated at least in part by the low-affinity FGF receptors (36 , 37) . In those instances in which the concentration of FGF molecules greatly exceeds the capacity of the low-and high-affinity receptors, steric hindrance may prevent receptor dimerization and consequent signaling.

We have shown that bFGF decreases Ewing’s sarcoma growth by inducing cell death. In vivo cell death was mediated, at least in part, by apoptosis. In vitro, electron microscopic features typical of both apoptosis and necrosis were identified. Although apoptosis and necrosis have been defined as two distinct modes of cell death, it is now clear that the two can occur simultaneously within a cell population and may involve common signaling and execution mechanisms. Populations of typically apoptotic and necrotic cells were identified in the bFGF-treated cell cultures, although cells with an intact nuclear membrane and chromatin margination, but with plasma membrane permeability, increased cytoplasmic volume and the loss of cellular organelles were also observed. These cells were scored as necrotic in the electron microscopic studies, although cell death in this cell population may better be described as oncosis, i.e., the development of necrosis accompanied by swelling and karyolysis (38) . FACS analysis of cells labeled with annexin V and PI seems to be a more useful quantitative method for the assessment of cell death, because it is not subjective, and comparisons between treated and nontreated cell populations can readily and rapidly be made. Our observations support the hypothesis that a single cytokine—in this case, bFGF—may simultaneously induce apoptosis and necrosis.

bFGF has previously been shown to decrease MCF-7 breast cancer cell proliferation (39, 40, 41) . Furthermore, bFGF and acidic FGF levels are lower in breast tumor biopsies than in normal breast tissue, which suggests that these factors may have an inhibitory role that is lost as tumors progress (42 , 43) . There has been some suggestion that a similar phenomenon may occur in SK-ES1 Ewing’s sarcoma cells (44) , although whether expression of bFGF correlates with outcome in tumors of the Ewing’s sarcoma family remains to be seen. These effects of bFGF are in direct contrast to its mitogenic and survival effects described in a number of different cell types of mesodermal and neuroectodermal origin (12) .

It is not known how bFGF stimulates growth in some tumors but inhibits growth in others. The effect of bFGF seems to be dependent on the growth phase of the cells investigated. Although bFGF inhibits apoptosis of oligodendrocytes under normal growth conditions, it can increase cell death when cells are prevented from entering the cell cycle (45) . Distinct patterns of FGF receptor expression and/or localization may confer such growth stimulatory or inhibitory effects. Although the Ewing’s sarcoma cell lines that were examined expressed all four of the high-affinity FGF receptors, a novel truncated form of FGF receptor-3 was identified that differed from the full-length receptor in its major ligand-binding region. It is possible that this may cause changes in structural conformation leading to constitutive activation of FGF receptor-3 signaling pathways such as STAT1 (46) , which results in growth inhibition and induction of cell death.

Because bFGF drives primitive pluripotent cells toward a differentiated neuronal phenotype (28) , we originally formed the hypothesis that treatment of Ewing’s sarcoma with bFGF and NGF might induce differentiation. However, we found no evidence of neural differentiation after treatment with bFGF alone, nor in combination with NGF. Furthermore, although studies in the neurally derived childhood tumor neuroblastoma have shown that induction of differentiation precedes apoptosis (47) , this does not seem to be the case in Ewing’s sarcoma. NGF was not mitogenic and did not offer a survival advantage for Ewing’s sarcoma cells in the presence or absence of bFGF, as reported in other neural cell types, including neuroblastoma (48 , 49) . This suggests bFGF signaling pathways are more important in Ewing’s sarcoma than those of NGF, as previously reported in pluripotent neural crest-derived stem cells (50) . The induction of cell death after exposure to bFGF may arise in Ewing’s sarcoma cells because the cells are unable to execute an appropriate differentiation response.

In summary, bFGF decreases the growth of Ewing’s sarcomas by inducing cell death. This may provide an opportunity for therapeutic initiatives. We are currently investigating the mechanism of bFGF-induced cell death, which may identify targets for the clinical modulation of Ewing’s sarcoma behavior.


   ACKNOWLEDGMENTS
 
We thank Carol Upton, Department of Electron Microscopy, Imperial Cancer Research Fund, Lincoln’s Inn Fields, London for assistance and advice on electron microscopy, and Del Watling and Sandra Peak, Biological Resources, Imperial Cancer Research Fund Clare Hall Laboratories, Potters Bar, Hertfordshire for assistance with in vivo mouse studies.


   FOOTNOTES
 
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.

1 Supported by the Candlelighter’s Trust, St. James’s University Hospital, Leeds, United Kingdom. Back

2 To whom requests for reprints should be addressed, at ICRF Cancer Medicine Research Unit, St. James’s University Hospital, Beckett Street, Leeds LS9 7TF, United Kingdom. Phone: 00-44-113-2065873; Fax: 00-44-113-2429886; E-mail: S.A.Burchill{at}leeds.ac.uk Back

3 The abbreviations used are: bFGF, basic FGF; FGF, fibroblast growth factor; BrdUrd, bromodeoxyuridine; TUNEL, terminal deoxynucleotidyl transferase-mediated nick end labeling; FACS, fluorescence-activated cell sorting; RT-PCR, reverse transcription-PCR; PI, propidium iodide; NGF, nerve growth factor. Back

Received 3/20/00. Accepted 9/ 1/00.


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 Western Blotting
 Immunofluorescence
 RT-PCR
 Statistical Analysis
 RESULTS
 DISCUSSION
 REFERENCES
 

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