The vascular endothelial growth factor (VEGF) receptor tyrosine kinase inhibitor sunitinib has been approved for first-line treatment of patients with metastatic renal cancer and is currently being trialled in other cancers. However, the effectiveness of this anti-angiogenic agent is limited by the presence of innate and acquired drug resistance. By screening a panel of candidate growth factors we identified fibroblast growth factor 2 (FGF2) as a potent regulator of endothelial cell sensitivity to sunitinib. We show that FGF2 supports endothelial proliferation and de novo tubule formation in the presence of sunitinib and that FGF2 can suppress sunitinib-induced retraction of tubules. Importantly, these effects of FGF2 were ablated by PD173074, a small molecule inhibitor of FGF receptor signalling. We also show that FGF2 can stimulate pro-angiogenic signalling pathways in endothelial cells despite the presence of sunitinib. Finally, analysis of clinical renal-cancer samples demonstrates that a large proportion of renal cancers strongly express FGF2. We suggest that therapeutic strategies designed to simultaneously target both VEGF and FGF2 signalling may prove more efficacious than sunitinib in renal cancer patients whose tumours express FGF2.
Tumour angiogenesis (the growth of new blood vessels into tumours) is a key driver of tumour progression and has received considerable attention as a therapeutic target (Hanahan and Folkman, 1996; Kerbel and Folkman, 2002; Ferrara and Kerbel, 2005). One of the most highly studied pro-angiogenic growth factors is vascular endothelial growth factor (VEGF), particularly the VEGF-A isoform, which is highly expressed in a variety of human tumours (Ellis and Hicklin, 2008). VEGF-A drives angiogenesis by signalling through two cognate receptors on local endothelial cells, VEGFR1 and VEGFR2 (Ferrara et al., 2003; Olsson et al., 2006). Binding of VEGF-A to VEGFR2 activates key intracellular signalling molecules including ERK1/2 and PLCγ. Pathways such as these coordinate the biological processes necessary for VEGF to induce the formation of new vessels (Olsson et al., 2006).
Numerous inhibitors of the VEGF signalling axis have been developed, including the VEGF-neutralising antibody, bevacizumab, and VEGF receptor tyrosine kinase inhibitors, such as sunitinib (Ferrara et al., 2004; Ferrara and Kerbel, 2005; Ellis and Hicklin, 2008; Kerbel, 2008). These drugs are particularly effective in VEGF-driven tumours such as clear-cell renal cancer. Indeed sunitinib monotherapy doubles progression-free survival and extends overall survival in this disease and is now standard first-line therapy (Motzer et al., 2007, 2009). Despite these impressive results, variation in response between individuals occurs and, more importantly, resistance to therapy invariably develops (Motzer et al., 2009; Rini and Atkins, 2009). Determinants of sunitinib response and the mechanisms that mediate the onset of resistance to sunitinib are poorly understood. This presents a significant barrier to the development of individualized therapy.
The tumour microenvironment contains numerous soluble factors, besides VEGF, that possess pro-angiogenic properties. These include growth factors that signal through tyrosine kinase receptors, such as members of the fibroblast growth factor (FGF), placental growth factor (PLGF) and platelet-derived growth factor (PDGF) families and factors that signal through non-tyrosine kinase receptors, such as interleukin-8 (IL-8) and transforming growth-factor-β (Relf et al., 1997; Ferrara, 2002; Presta et al., 2005). These factors may have an important role in the induction of tumour angiogenesis and may also be involved in altering the responsiveness of tumour blood vessels to inhibitors of the VEGF signalling axis (Bergers and Hanahan, 2008). In support of this latter concept, FGF2 is upregulated in a mouse tumour model of acquired resistance to DC101 (a VEGFR2 inhibitory antibody) and inhibition of FGF2 in combination with DC101 led to improved anti-tumour responses (Casanovas et al., 2005). Moreover, inhibition of PLGF or PDGF has been reported to enhance the anti-tumour efficacy of VEGF pathway inhibitors in some murine tumour models (Bergers et al., 2003; Fischer et al., 2007; Crawford et al., 2009). Another recent study demonstrated that IL-8 is upregulated in sunitinib-resistant tumour xenografts and that inhibition of IL-8 can re-sensitize these resistant tumours to sunitinib (Huang et al., 2010b).
Sunitinib is an orally available tyrosine kinase inhibitor that potently inhibits several angiogenesis-related tyrosine kinase receptors that is, VEGF receptors (VEGFR1, VEGFR2 and VEGFR3), PDGF receptors (PDGFRα and PDGFRβ), and also the KIT receptor (Abrams et al., 2003; Mendel et al., 2003; O’Farrell et al., 2003). A recent study that examined the activity of sunitinib against a panel of 317 kinases (representing >50% of the known kinome) revealed that sunitinib has relatively high affinity for at least 40 other kinases, including myosin light-chain kinase, MEK1 and MEK2 (Karaman et al., 2008). As activation of multiple kinase-signalling pathways is a requirement for growth factor-mediated stimulation of angiogenesis, the relatively broad kinase substrate specificity of sunitinib might be expected to limit the potential for other growth factors to drive resistance to sunitinib through a mechanism that involves direct stimulation of endothelial cells.
Here we identify FGF2 to be a potent mediator of endothelial cell resistance to sunitinib. We further demonstrate that FGF2 can suppress the anti-angiogenic activity of sunitinib by directly stimulating pro-angiogenic signalling in endothelial cells. We also show that FGF2 is expressed in a subset of human renal cancers. These data provide strong support for simultaneous targeting of VEGF and FGF signalling as a therapeutic strategy in renal cancer.
A screen of pro-angiogenic growth factors identifies FGF2 as a mediator of endothelial cell resistance to sunitinib
Sunitinib is a potent inhibitor of VEGFR2 tyrosine kinase activity. In agreement with previous reports (Mendel et al., 2003), nanomolar concentrations (10–100 nM) of sunitinib inhibited VEGF-induced phosphorylation of VEGFR2 in human endothelial cells (Figure 1a) and VEGF-induced endothelial cell proliferation (Figure 1b). These concentrations are clinically relevant, as circulating concentrations of sunitinib measured in patients are in the nanomolar range and do not typically exceed 200 nM (Faivre et al., 2006; Britten et al., 2008).
To identify growth factors potentially involved in resistance to sunitinib, we screened a panel of 19 pro-angiogenic growth factors for their ability to promote endothelial proliferation in cell cultures incubated with VEGF and 100 nM sunitinib (Figure 1c). Although most of the growth factors tested were able to induce a small degree of endothelial cell proliferation in the presence of sunitinib, only FGF1 and FGF2 were able to restore endothelial cell proliferation to levels that matched (FGF1) or exceeded (FGF2) those observed in the absence of sunitinib (Figure 1c). These data therefore identify FGF1, and more strikingly FGF2, as candidate factors that may be potent mediators of endothelial cell resistance to sunitinib.
FGF2 suppresses the anti-angiogenic activity of sunitinib in a 3-dimensional assay of endothelial tube formation
As FGF2 was identified as the most potent factor able to drive resistance to sunitinib in an endothelial cell proliferation assay (Figure 1c), we proceeded to validate this activity within an in vitro model of angiogenesis. Human umbilical vein endothelial cells (HUVEC)-coated beads were embedded in a fibrinogen gel and then incubated with VEGF and different concentrations of sunitinib. After 7 days, endothelial tubules had sprouted from VEGF alone-treated beads, but tubule formation was significantly inhibited by low concentrations (10–100 nM) of sunitinib (Figures 2a and b) confirming that sunitinib can inhibit VEGF-stimulated tubule formation in this assay. Importantly, when the assay was repeated in the presence of VEGF and FGF2, the number of tubules was enhanced 3–4 fold compared with beads cultured with VEGF alone (Figure 2c). These data are in agreement with previous reports showing that FGF2 can enhance VEGF-mediated angiogenesis (Pepper et al., 1992; Giavazzi et al., 2003). Sunitinib was also able to suppress tubule formation in the presence of VEGF and FGF2 (Figure 2c). However, tubule formation in the presence of VEGF, FGF2 and sunitinib was still significantly enhanced compared with tubule formation in the presence of VEGF and sunitinib (Figure 2c). These data confirm that FGF2 can act to suppress the anti-angiogenic effects of sunitinib.
Having shown that FGF2 can suppress the effects of sunitinib on de novo tubule formation, we then addressed whether FGF2 could suppress the effects of sunitinib on pre-formed tubules. HUVEC-coated beads embedded in a fibrinogen gel were incubated from the start of the assay with VEGF in the presence or absence of FGF2. After 7 days, when tubules were clearly visible, the medium was then additionally supplemented with either vehicle or 10 nM sunitinib. The behaviour of individual tubules was monitored by time-lapse microscopy for the following 48 h. In cultures incubated with VEGF and vehicle, tubules grew in length through extension at the tip of the tubule (Figure 2d and Supplementary Movie S1). In contrast, in cultures incubated with VEGF and 10 nM sunitinib, tubules rapidly retracted (Figure 2e and Supplementary Movie S2). These data are consistent with previously published data showing that inhibitors of VEGF receptor tyrosine kinase activity can induce the retraction of existing vessels (Mancuso et al., 2006). However, in the combined presence of VEGF, FGF2 and 10 nM sunitinib, tubule retraction was abrogated (Figure 3f and Supplementary Movie S3). We quantified the ability of FGF2 to suppress tubule retraction, by measuring the change in length of tubules after addition of 10, 25, 50 or 100 nM sunitinib. In the presence of VEGF alone, sunitinib induced tubule retraction in a dose-dependent manner (Figure 2g). However, in the presence of VEGF and FGF2, the ability of sunitinib to induce tubule retraction was significantly suppressed (Figure 2g). These data show that low concentrations of sunitinib (10–100 nM) can induce the retraction of pre-formed tubules, but that this effect is suppressed by FGF2.
PD173074 inhibits FGF2-mediated resistance to sunitinib
We then tested whether the effects of FGF2 on endothelial cell proliferation could be suppressed using an inhibitor of FGF receptor signalling. The small molecule tyrosine kinase inhibitor PD173074 is a potent inhibitor of FGF receptor tyrosine kinase activity (Mohammadi et al., 1998; Pardo et al., 2009). We tested the relative ability of sunitinib and PD173074 to inhibit VEGF- and FGF2-mediated endothelial cell proliferation. Inhibition of endothelial cell proliferation was assessed in the presence of 100 ng/ml VEGF or in the presence of 2.5 ng/ml FGF2, which is an FGF2 concentration that induces equivalent levels of endothelial proliferation to 100 ng/ml VEGF in our assay. Sunitinib concentrations in the range of 10–100 nM inhibited VEGF-stimulated proliferation of endothelial cells, but did not inhibit FGF2-stimulated proliferation of endothelial cells (Figure 3a). Conversely, PD173074 concentrations in the range of 10–100 nM inhibited FGF-stimulated proliferation of endothelial cells, but did not inhibit VEGF-stimulated proliferation of endothelial cells (Figure 3b). These data confirm the ability of sunitinib to selectively inhibit VEGF-mediated endothelial cell proliferation and the ability of PD173074 to selectively inhibit FGF2-mediated endothelial cell proliferation in this assay.
We then tested whether PD173074 could restore sunitinib sensitivity to endothelial cells cultured in the presence of VEGF and FGF2. The combination of VEGF and FGF2 resulted in enhanced proliferation of endothelial cells compared with cells cultured with VEGF alone and also reduced the sensitivity of cells to sunitinib (Figure 3c). However, addition of 50 nM PD173074 to cells cultured with FGF2 and VEGF prevented the FGF2-mediated enhancement in endothelial cell proliferation and restored sensitivity to sunitinib (Figure 3c). The effects of PD173074 on tubule retraction were also examined. As observed previously, the presence of FGF2 inhibited the retraction of pre-formed tubules (Figure 3d). However, the addition of 50 nM PD173074 abrogated this effect of FGF2 and restored sunitinib sensitivity in this assay (Figure 3d). These data show that an inhibitor of FGF receptor signalling can prevent FGF2 from suppressing the sensitivity of endothelial cells to sunitinib.
FGF2 activates pro-angiogenic signalling pathways in endothelial cells in the presence of sunitinib
Having confirmed that FGF2 can suppress the anti-angiogenic activity of sunitinib, we then addressed whether FGF2 can activate pro-angiogenic signalling pathways in endothelial cells in the presence of sunitinib. Stimulation of HUVECs with VEGF alone promoted strong phosphorylation of MEK1/2 and ERK1/2, but this was suppressed by sunitinib (Figure 4). In contrast, in the presence of FGF2 alone or FGF2 and VEGF combined, strong phosphorylation of MEK1/2 and ERK1/2 occurred in both the presence and absence of sunitinib (Figure 4). Stimulation of HUVECs with VEGF alone also promoted phosphorylation of PLCγ, which again was suppressed by sunitinib (Figure 4). Addition of FGF2 alone to HUVECs also promoted phosphorylation of PLCγ, although activation of this pathway appeared to be weaker than that observed with VEGF. However, the weak activation of PLCγ by FGF2 was not blocked by sunitinib. These data show that although sunitinib can inhibit VEGF-mediated activation of ERK1/2 and PLCγ, sunitinib does not block FGF2-mediated activation of ERK1/2 and PLCγ.
FGF2 expression in human renal cancer samples
Sunitinib has been approved for the first-line treatment of patients with metastatic renal cancer. Renal cancers are highly angiogenic and sunitinib is proposed to suppress the growth of these tumours mainly through inhibition of angiogenesis (Faivre et al., 2007; Huang et al., 2010a). As our data identified FGF2 as a potent suppressor of sunitinib's anti-angiogenic activity, we assessed the expression of FGF2 in human renal cancer samples. An FGF2-specific antibody was used to stain a tissue microarray (TMA) composed of renal cancer samples from a cohort of 74 patients with advanced renal cancer who had progressed on conventional cytokine therapy, but who did not receive sunitinib therapy (more detailed information on these patients can be found in Table 1 and the Materials and methods section). Interestingly, we found that FGF2 staining was prominent in both tumour cells and in tumour blood vessels (Figures 5a–d and Table 2). Tumour-cell expression of FGF2 was localized predominantly to the nuclei of tumour cells (for an example see Figure 5a) and was detectable in 36 of the 74 cases examined and absent from the remaining 38 cases (for an example of negative staining see Figure 5b). Staining of the tumour vasculature was localized to the cytoplasm of endothelial cells or associated with the vascular basement membrane (for an example see Figure 5c) and was detected in 41 of the 74 cases examined, but was absent from the remaining 33 cases (for an example of negative staining see Figure 5d). These data confirm previous work showing that FGF2 is expressed in human renal cancers (Eguchi et al., 1992; Nanus et al., 1993; Slaton et al., 2001; Fukata et al., 2005). Moreover, we clearly show that FGF2 expression in advanced renal cancer can be associated with the tumour compartment or the vascular compartment.
Resistance to therapy is a major limiting factor in the successful treatment of cancers with anti-angiogenic agents that target the VEGF signalling axis (Bergers and Hanahan, 2008; Ebos et al., 2009). Stimulation of tumour angiogenesis by other pro-angiogenic growth factors in the tumour microenvironment may be one mechanism through which this resistance occurs. Previous work has shown that inhibition of FGF2 or PLGF can overcome acquired resistance to VEGFR2 inhibition in mouse tumour models (Casanovas et al., 2005; Fischer et al., 2007). However, to the best of our knowledge, the direct ability of growth factors to stimulate angiogenesis in human endothelial cells when VEGFR2 is inhibited by sunitinib has not been clearly demonstrated. Here we screened a panel of 19 pro-angiogenic factors and revealed that FGF2 can potently suppress the anti-angiogenic activity of sunitinib. We show that FGF2 is able to directly stimulate endothelial cell proliferation and endothelial tubule formation in the presence of sunitinib. We also show that FGF2 can suppress sunitinib-induced retraction of pre-formed endothelial cell tubules. These data suggest that the pro-angiogenic function of FGF2 may be directly relevant for resistance to sunitinib.
It was surprising to find that the other pro-angiogenic factors we tested showed significantly less activity in comparison with FGF2. This may reflect a limitation of our in vitro screening approach and we cannot exclude the possibility that some of the angiogenic factors screened may indeed have a role in resistance to sunitinib in vivo. Of note, a recent study demonstrated a role for IL-8 in mediating resistance to sunitinib in vivo (Huang et al., 2010b), whereas our screen did not reveal a role for IL-8. It is likely that tumour-specific microenvironmental effects, which are not recapitulated in our assay, may permit certain growth factors to influence the response of endothelial cells to sunitinib in vivo. Moreover, some growth factors may mediate resistance to sunitinib by acting on other cell types, such as pericytes, tumour cells or infiltrating immune cells. Therefore, complementary work using appropriate in vivo models of resistance to sunitinib may demonstrate a role for other growth factors in mediating tumour resistance to sunitinib.
It is known that VEGF and FGF2 can activate similar signalling pathways in endothelial cells in vitro, such as the Ras–Raf–MEK–ERK1/2 pathway and the PLCγ–PKC pathway (Cross and Claesson-Welsh, 2001; Presta et al., 2005; Olsson et al., 2006). Moreover, essential roles for both of these pathways in the process of angiogenesis have been demonstrated (Presta et al., 1991; Eliceiri et al., 1998; Meyer et al., 2003; Mavria et al., 2006). Here we show that although sunitinib can inhibit VEGFR2-mediated activation of ERK1/2 and PLCγ, sunitinib did not prevent FGF2-mediated activation of ERK1/2 and PLCγ. These data suggest that FGF2 can bypass inhibition of VEGFR2 signalling by activating at least two key pro-angiogenic signalling pathways. It is likely that FGF2-mediated angiogenesis also involves activation of other signaling pathways that we have not examined here. Although we do not suggest that FGF2-mediated activation of ERK1/2 and PLCγ is sufficient for FGF2 to activate angiogenesis in the presence of sunitinib, these data demonstrate the principle that pro-angiogenic signalling in response to FGF2 can take place in endothelial cells in the presence of sunitinib.
We examined the expression of FGF2 in tumour samples obtained from metastatic renal cancer patients. As well as confirming that FGF2 is commonly expressed in these tumours, we found that FGF2 can be differentially localized in patient samples. Expression of FGF2 in tumour nuclei was observed in 49% of the samples, whereas expression of FGF2 in tumour blood vessels could be demonstrated in 55% of the samples. Expression of FGF2 in the nuclei of tumour cells in human cancer specimens has been reported in other studies (Eguchi et al., 1992; Joy et al., 1997) and functional studies have shown that fibroblast growth factors can have a signalling role in the nucleus (Bryant and Stow, 2005). The origins of the vascular FGF2 staining we observed are currently unclear. As tumours can secrete FGF2, this staining pattern may conceivably be due to the accumulation of secreted FGF2 in and around tumour blood vessels. However, 26% of the cases we examined showed positive FGF2 staining in the vasculature, whereas the tumour cells in the same sample were negative for FGF2 staining (see Table 2). Therefore, it is possible that the vascular FGF2 staining we observed may represent selective upregulation of FGF2 expression in the tumour endothelium. In support of this, expression of FGF2 by endothelial cells and autocrine signalling of FGF2 in endothelial cells has been previously described (Schweigerer et al., 1987; Gualandris et al., 1996; Pintucci et al., 2002). It will now be important to examine whether the expression and localisation of FGF2 in renal cancers has any role in determining the response of renal cancer patients to sunitinib. If tumour or vascular expression of FGF2 in renal cancer has a negative impact on response to sunitinib, then this could represent a potential biomarker to identify renal cancer patients who are unlikely to respond to sunitinib. These patients might then be considered for treatment with one of the alternative targeted agents currently being tested in renal cancer, such as novel agents that are designed to inhibit both VEGF receptor and FGF receptor signalling.
Several kinase inhibitors which target both VEGF and FGF receptor signalling are currently being trialled in patients (Turner and Grose, 2010). Although our data suggest that this may be an attractive therapeutic approach in renal cancer, it remains to be seen whether targeting both of these pathways in renal cancer will be more efficacious than targeting the VEGF pathway alone. The efficacy of this approach will likely depend on several factors, including (1) the relative importance of VEGF and FGF2 signalling as oncogenic drivers in a particular tumour, (2) the presence of alternative resistance mechanisms in a particular tumour and (3) the level of toxicity experienced by patients treated with these novel-kinase inhibitors.
In conclusion, our data suggest that the pro-angiogenic function of FGF2 may be relevant for resistance to sunitinib. This may be especially important in renal cancer, in which FGF2 expression is prominent. We suggest that therapeutic strategies designed to simultaneously target both VEGF and FGF2 signalling may prove more efficacious than sunitinib in renal cancer patients whose tumours express FGF2.
Materials and methods
Cell culture reagents were obtained from the following sources: fetal calf serum (FCS) (Gibco, Invitrogen Ltd, Paisley, UK) and endothelial cell mitogen from bovine brain (Serotech, Kidlington, Oxfordshire, UK). Antibodies were obtained from the following sources: total MEK (BD Pharmingen, San Jose, CA, USA), pThr202/pTyr201-ERK1/2, pSer217/pSer221-MEK, pTyr783-PLCγ, PLCγ, pTyr1175-VEGFR2, VEGFR2 (Cell Signalling Technology, Danvers, MA, USA), FGF2 (Peprotech, London, UK), HSC-70 (Santa Cruz Biotechnology, Santa Cruz, CA, USA). All growth factors utilised were obtained from Peprotech, except for angiopoietin 2 and erythropoietin that were obtained from R&D Systems (Abingdon, Oxfordshire, UK). Sunitinib malate was obtained from LC laboratories (Woburn, MA, USA) and PD173074 was obtained from Sigma (Poole, Dorset, UK). Unless otherwise stated, all other reagents were obtained from Sigma.
HUVECs from pooled donors (TCS Cell Works, Buckingham, Buckinghamshire, UK) were cultured in HUVEC-specific medium (M199 medium supplemented with 20% FCS, 20 μg/ml endothelial cell mitogen, 10 μg/ml heparin and antibiotics). Cells were used for experiments at passage 4–8. Normal human dermal fibroblast cells (TCS Cell works) were cultured in Dulbecco's Modified Eagle Medium supplemented with 10% FCS and antibiotics.
HUVEC proliferation assays
HUVECs were plated on 96-well plates at a density of 1000 cells per well in HUVEC-specific medium. The next day the medium was changed for M199 plus 10% FCS, supplemented with 100 ng/ml VEGF and with sunitinib or PD173074 at the indicated concentration or vehicle (0.1% DMSO; dimethyl sulfoxide). For the purpose of the growth factor screen, cells were incubated with M199 medium plus 10% FCS, supplemented with 100 ng/ml VEGF, 50 ng/ml of an alternative growth factor and 100 nM sunitinib. After 72 h, cell viability was quantified using the Cell Titre Glow cell-viability reagent (Promega, Southampton, Hampshire, UK), according to the manufacturer's instructions. The plates were read in a luminescence plate reader (PerkinElmer, Cambridge, Cambridgeshire, UK).
HUVECs were plated on 100 mm diameter plates (at a density of 5 × 105 per plate) and cultured for 48 h. The cells were washed three times in serum-free M199 medium and then starved for 3 h in serum-free M199 medium. Treatment with the indicated concentration of sunitinib was applied 15 min before stimulation with the indicated concentrations of VEGF and/or FGF2. After 5 or 10 min, plates were transferred to ice, washed twice with ice-cold phosphate-buffered saline and then protein lysates were collected in lysis buffer (1% IGEPAL CA-630, 20 mM Tris pH 7.5, 150 mM NaCl, 10% glycerol, 1 mM Na3VO4, 10 mM NaF, 1 mM 4-(2-aminoethyl) benzynesulphonyl fluoride, 0.8 mM aprotinin, 0.05 mM bestatin, 0.015 mM E-64, 0.02 mM leupeptin and 0.01 mM pepstatin). Cell lysates were mixed with reducing Laemmli sample buffer, separated on 7% or 10% SDS–polyacrylamide gels and transferred onto Hybond–ECL nitrocelluose membranes (GE Healthcare Life Sciences, Little Chalfont, Buckinghamshire, UK) over night. Membranes were blocked at room temperature for 1 h in TBS-T (100 mM Tris–HCl, 0.2 M NaCl, 0.1% Tween 20 v/v) containing 5% milk (Marvel, Premier International Food, Spalding, Lincolnshire, UK) followed by primary antibody incubation for 1 h at room temperature or overnight at 4°C in TBS-T containing 5% bovine serum albumin. After washing, incubation with horse radish peroxidase-conjugated secondary antibodies was performed for 1 h at room temperature in TBS-T containing 5% milk. Signal was visualized with the enhanced chemiluminescence reagent (GE Healthcare Life Sciences).
Endothelial tube formation assays
Tube formation assays were performed using a modified version of a previously published protocol (Nakatsu and Hughes, 2008). In brief, HUVECs were cultured for 24 h in EGM2 complete medium (Lonza, Slough, Berkshire, UK) and then cells were trypsinised and resuspended in EGM2 at a concentration of 2 × 106 HUVECs per ml. Cytodex3 beads (GE healthcare Life Sciences) were coated with HUVECs by incubating 2 × 106 HUVECs with 3 × 104 beads for 4 h with gentle agitation every 20 min. Coated beads were then diluted to a volume of 5 ml in EGM2 and placed in a tissue-culture incubator overnight. The next day, beads were washed in EGM2 and resuspended in 50 ml of sterile 2 mg/ml fibrinogen in phosphate-buffered saline. A volume of 500 μl of bead-fibrinogen solution was then transferred to each well of a 24-well plate (approximately 100 beads per well) containing 6.25 μl 50 U/ml thrombin. Fibrinogen gels were allowed to set for 15 min at 37°C and then 2 × 104 fibroblasts in EGM2 were seeded on top of each well. After 3 h, medium was exchanged for EBM2 (Lonza) plus 2.5% FCS supplemented with growth factors (10 ng/ml VEGF and/or 50 ng/ml FGF2), sunitinib, PD173074 or vehicle (0.1% DMSO) as indicated. The plates were re-fed every 2 days.
Analysis of tube formation assays
The quantity of tubules that sprouted from HUVEC beads was measured after 7 days of culture by fixing beads in 4% formalin and then counting the number of sprout tips per bead under an inverted light microscope. A total of 15 beads were counted per well and all conditions were counted in duplicate wells. For time-lapse microscopy, tubule formation assays were established as described above and after 7 days the medium was supplemented with sunitinib or vehicle (0.1% DMSO) as indicated. The plates were then transferred to the stage of an inverted Leica IX-70 microscope fitted with a heated chamber and a supply of 10% CO2. Images were captured every 20 min for a total of 48 h. For the quantification of tubule extension/retraction, cultures were incubated with VEGF alone or VEGF and FGF2, as indicated, for 7 days in an incubator with 37°C/10% CO2. Plates were then transferred to the stage of the inverted Leica IX-70 microscope and photomicrographs were captured of individual tubules (time point=0 h). The medium was then supplemented with sunitinib, PD173074 or vehicle (0.1% DMSO), as indicated, and plates returned to the incubator. After 48 h, photomicrographs of the same tubules were recorded again (time point=48 h). Sprout length was quantified from the photomicrographs recorded at 0 and 48 h by using image analysis software (Adobe Photoshop, Uxbridge, UK). Percentage change in tubule length was calculated using the formula: 100 × (a/b)−1, where a is the tubule length at 48 h and b the tubule length at 0 h.
Renal cancer samples were collected from patients with advanced renal cancer who were enroled in a Phase III clinical trial of lapatinib versus hormone therapy (Ravaud et al., 2008). Tissues were collected from patients before commencing treatment. The tissue was fixed in formalin, embedded in paraffin and used to generate a TMA consisting of three cores from each patient sample. For the purpose of this study, we only analysed renal cancer samples of confirmed clear cell histology that were collected from 74 patients in the hormone therapy arm of the trial (patients received daily megesterol acetate or daily tamoxifen until disease progression or withdrawal from the trial). Following de-waxing and processing for antigen retrieval, the TMA was stained with a rabbit anti-human FGF2 antibody (Peprotech) that was then detected using horse radish peroxidase-conjugated secondary antibodies and DAB staining. Scoring of the TMA was performed by two observers: Daniel Berney (a genitourinary pathologist) and Andrew Reynolds (the lead investigator). Scoring was recorded separately for FGF2 expression in tumour nuclei and FGF2 expression in the vasculature. Individual patient cases were scored as positive if one or more of the three cores from each patient stained positive for nuclear FGF2 or vascular FGF2.
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We would like to thank Alan Ashworth, Clare Isacke and Nicholas Turner for critical comments on the paper and Breakthrough Breast Cancer for research funding.
Thomas Powles is the recipient of an educational research grant from Pfizer Global Pharmaceuticals. The other authors declare no potential conflict of interest.
Supplementary Information accompanies the paper on the Oncogene website
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Welti, J., Gourlaouen, M., Powles, T. et al. Fibroblast growth factor 2 regulates endothelial cell sensitivity to sunitinib. Oncogene 30, 1183–1193 (2011). https://doi.org/10.1038/onc.2010.503
- growth factors
- renal cancer
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