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Molecular Biology, Pathobiology and Genetics |
1 Molecular Oncology Laboratories, Cancer Research UK, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital; 2 Department of Urology, Churchill Hospital, University of Oxford, Oxford; and 3 In Situ Hybridization Service, Cancer Research UK, London, United Kingdom
Requests for reprints: Ji-Liang Li, Cancer Research UK, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Oxford OX3 9DS, United Kingdom. Phone: 44-1865-222457; Fax: 44-1865-222431; E-mail: lij{at}cancer.org.uk or aharris.lab{at}cancer.org.uk.
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Abstract |
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. Down-regulation of DLL4 expression with RNA interference led to decreased expression of HEY1 and EphrinB2, and the inhibition of endothelial cell proliferation, migration, and network formation, all of which are important processes in tumor angiogenesis. The inhibition of proliferation occurred via the induction of cell cycle arrest in G0-G1 by increased expression of p21 and decreased phosphorylation of retinoblastoma. We conclude that an optimal window of the DLL4 expression is essential for tumor angiogenesis and that selective modulation of the DLL4 expression within human tumors may represent a potential novel antiangiogenic therapy. |
Introduction |
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and HIF-2 in response to hypoxia leads to the up-regulation of numerous adaptive proteins, including angiogenic factors such as vascular endothelial growth factor (VEGF; ref. 2).
Many of the processes involved in tumor angiogenesis are mirrored during embryonic vascular development. In the past decade, the Notch signaling pathway has been shown to play a key role in vascular development. The Notch pathway is an evolutionary conserved intercellular signaling pathway involved in numerous biological processes including cell fate determination, cellular differentiation, proliferation, survival, and apoptosis (3–5). In mammalian cells, there are five transmembrane Notch ligands [Jagged1, Jagged2, delta-like 1 ligand (DLL1), DLL3, and DLL4] and four Notch receptors (Notch1-4). Ligand-receptor binding activates the signaling pathway by initiating the cleavage of the Notch intracellular domain from the cell membrane (6, 7). The Notch intracellular domain subsequently translocates to the nucleus where it classically interacts with the transcription factor C promoter binding factor 1 (CBF1)/recombination signal binding protein J
(RBP-J)/Suppressor of Hairless [Su(H)]/Lag-1 protein (CSL). The basic helix-loop-helix-orange proteins (bHLH) Hairy/Enhancer of Split (HES1, 5, and 7) and the HES-related proteins (HEY1, HEY2, and HEYL) are the best-characterized downstream targets of the Notch intracellular domain/CSL complex (8–10).
Mice deficient for various Notch components die in utero of severe vascular abnormalities (11). DLL4 knockout mice display vascular defects similar in pattern to those seen in Notch1/Notch4 double knockouts. Interestingly, haploinsufficiency of DLL4 also results in embryonic lethality from severe vascular defects (12–14). Haploinsufficiency for angiogenic factors is uncommon and highlights the importance of DLL4 over and above other components of the Notch pathway. Similar phenotypes have been only described for VEGF knockout mice (15, 16).
We and others have previously shown that DLL4 is expressed at sites of vascular development and angiogenesis (14, 17, 18). During early embryonic development, DLL4 is initially observed in major arterial vessels; with embryonic maturation, this pattern of expression disappears whereupon it is confined to small vessels and capillaries. DLL4 is up-regulated at sites of physiologic and pathologic angiogenesis (17). It is high in ovarian vessels surrounding growing follicles and is up-regulated within the tumor vasculature when compared with adjacent normal tissues. Xenograft studies showed high expression of mouse DLL4, but not of human-derived DLL4, within the tumor vasculature. We have shown in vitro that DLL4 is a hypoxia-regulated gene, although the pathway was unknown (17).
In this study, we investigated the expression of DLL4 in renal tumors, analyzed the relationship between DLL4, hypoxia, and angiogenic growth factors, and assessed the biological function of DLL4 in tumor angiogenesis.
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Materials and Methods |
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RNA interference. Chemically synthesized gene-specific small interfering RNA (siRNA) duplexes were designed, BLAST searched, and manufactured by Eurogentec (Seraing, Belgium) as per accepted guidelines (20). The following siRNA sequences were used: DLL4-Duplex1 (20 nmol/L): CUACUAUGGAGACAACUGCTT; DLL4-Duplex2 (20 nmol/L): UGACCACUUCGGCCACUAUTT; DLL4 scrambled control (20 nmol/L): UCUGAAAAGCACGCUUGACTT; HIF-1 (50 nmol/L): CUGAUGACCAGCAACUUGATT; HIF-2 (50 nmol/L): CAGCAUCUUUGAUAGCAGUTT; and HIF scrambled control (50 nmol/L): AGUUCAACGACCAGUAGUCTT. Transfection of siRNA duplexes was done with cells at 40% to 50% confluency using 3 mL of Optimem-1 and 12 µL of Lipofectamine 2000 (Invitrogen, Carlsbad, CA). Biological functional assays and gene expression analysis were done 24 hours after siRNA transfection.
RNA extraction and reverse transcription. Extraction of total RNA from cultured endothelial cells was done using TRI-Reagent (Sigma-Aldrich) as per instructions of the manufacturer. The quality of RNA extracted was assessed using RNA 6000 Nano Chips and the Agilent 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA). cDNA was synthesized by reverse transcribing total RNA with the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA).
Real-time quantitative PCR. Real-time quantitative PCR reactions were done in triplicate using the Corbett Research Rotor Gene RG-3000 (Sydney, Australia). Each reaction was done in an individual tube and made to 25 µL containing the equivalent of 5 ng reverse-transcribed RNA (1 ng in the case of 18S rRNA), 12.5 µL TaqMan 2x PCR Master Mix (Applied Biosystems), and 1.25 µL the probe/primer mix. Human ß-actin (renal samples) and eukaryotic 18S rRNA (endothelial cells) were used as reference genes to normalize for differences in the amount of total RNA in each sample. The following primer/probe kits were purchased as Assays-on-Demand from Applied Biosystems: DLL4 (Hs00184092_m1), HES1 (Hs00172878_m1), HEY1 (Hs00172878_m1), HEY2 (Hs00232622_m1), Ephrin B2 (Hs00187950_m1), VEGFR2 (Hs00176676_m1), CD34 (Hs00156373_m1), and VEGF (Hs00173626_m1). The reaction efficiency for each gene was calculated after obtaining standard curves for each PCR reaction by making 2-fold serial dilutions covering the range equivalent to 20 to 0.625 ng RNA (5-0.125 ng for 18S rRNA). Relative quantitation of gene expression was done using the method described by Pfaffl (21). The relative ratio of gene expression was calculated as [(Etarget)
CtTARGET(mean comparator – mean sample)] ÷ [(Eref)CtREF(mean comparator – mean sample)], where Etarget is reaction efficiency of the gene of interest; Eref, reaction efficiency of the reference gene; and Ct, the cycle difference between the comparator and the sample. All calculations are based on the mean value of PCR reactions done in triplicate. The comparator for the clinical samples was the median normal kidney; the comparator for cell line work is stated accordingly in the figure legends.
Reverse transcription-PCR. Standard PCR reactions were done using HotStarTaq DNA polymerase (Qiagen, Valencia, CA). In brief, 2x HotStarTaq master mix was combined with 1 µmol/L primers and 5 ng cDNA in a total volume of 20 µL. PCR cycling was done for 38 cycles. PCR products were separated on 1.7% agarose gels stained with ethidium bromide. The following primer sequences were used:
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For the matrigel network formation assay, each well of a 24-well Falcon tissue culture plate was evenly coated with 250 µL Matrigel (BD Biosciences, Bedford, MA). 4 x 104 HUVECs were seeded 24 hours posttransfection in triplicate per well in MCDB131 + 20% FCS + ECGS. Network formation was assessed after 16 hours by photographing the matrices using a Zeiss Axiovet 135 inverted light microscope and a Nikon CoolPix 4500 digital camera. Quantification of tube formation was done using Adobe Photoshop 7.0.
For the cell migration assays, following transfection and quiescing for 24 hours, 5 x 104 cells were seeded in triplicate in MCBD131 + 1% FCS into the insert well of the BD BioCoat Endothelial Cell Migration Assay plate (BD Biosciences; ref. 24). Migration across the microporous membrane occurred in response to stimuli within the lower chamber (MCDB131 + 5% FCS, MCBD131 + 1% FCS + 10 ng/mL VEGF165; R&D Systems, Minneapolis, MN). Following a 22-hour incubation, the cells were labeled with Calcein AM (Molecular Probes, Eugene, OR) and the fluorescent signal from the undersurface of the membrane measured using a BMG FLUOstar fluorescence plate reader. Images of the insert well were captured using a Zeiss Axiovet 135 inverted light microscope and an Axiocam digital camera.
Scratch wound assay was done as described (25). Twenty-four hours after transfection, 4 x 105 HUVECs were seeded in triplicate into a six-well Falcon tissue culture plate. Cells were fluorescently labeled with 5 µmol/L CMFDA CellTracker dye (Molecular Probes); after which, the monolayer was "scratched" and incubated in MCDB131 + 10% FCS + 50 ng/mL VEGF + 2.5 µg/mL Mitomycin C (Sigma-Aldrich). The wounded area was photographed using a Zeiss Axiovet 135 inverted light microscope and a Nikon CoolPix 4500 digital camera. Quantitation of wound width was done using Adobe Photoshop 7.0.
Cell cycle analysis. Twenty-four hours after transfection, synchronization of cell cycles was achieved by culturing HUVECs for 24 hours in MCDB131 + 2% FCS. Cells were subsequently cultured for 24 hours in either MCDB131 + 20% FCS + ECGS or MCDB131 + 2% FCS, and then fixed in 70% ethanol. Cell cycle analysis was done by fluorescence-activated cell sorting analysis after pretreating cells with RNase (100 µg/mL) and staining nuclear DNA with propidium iodide (50 µg/mL, Sigma-Aldrich; ref. 23).
Apoptosis analysis. 1 x 105 HUVECs were seeded in triplicate in MCDB131 + 20% FCS + ECGS into a 24-well Falcon tissue culture plate. After 24 hours, the cells were serum and growth factor deprived using MCDB131 + 1% FCS. Cell viability was assessed by periodically counting the number of adherent cells using the Beckman Z2 Coulter counter. Apoptosis was assessed using the Apo-ONE Homogenous Caspase 3/7 Assay (Promega Corp., Madison, WI; ref. 26). 2 x 104 HUVECs were seeded in triplicate in 100 µL MCDB131 + 20% FCS + ECGS with 100 µL Apo-ONE reagent in a 96-well white-walled tissue culture plate. After 4 hours, fluorescence was measured at 485/530 nm using a Bio-Tek Flx800 fluorescence plate reader.
In situ hybridization. Radioactive in situ hybridization on paraffin-embedded sections was done by the in situ hybridization service, Cancer Research UK (London), using a method we described previously (17). The probe used in the in situ hybridization for DLL4 was a 741 bp fragment located from position 1,775 to 2,516 bp (17).
Western blotting. Protein extraction and Western blotting were done as previously described (27). Antibodies against the following proteins were used for detection: HIF-1 (1:1,000; BD PharMingen, San Diego, CA), HIF-2 (1:1,000; Abcam, Cambridge, MA), p21 (1:100; Calbiochem, San Diego, CA), p27 (1:500; Cell Signaling), retinoblastoma (1:500; BD PharMingen, Bedford, MA), and ß-tubulin (1:10,000; Sigma-Aldrich).
Immunohistochemistry. Immunohistochemistry was done as previously described (28). In brief, paraffin-embedded sections were dewaxed, rehydrated, and incubated with the anti-CD34 monoclonal antibody QBEnd 10 (Novocastra, Newcastle, United Kingdom). Visualization was achieved using the EnVision HRP kit (DAKO Cytomation, Carpinteria, CA).
Statistics. Statistical analysis was done using GraphPad Prism. Statistical tests used included t test, one-way ANOVA, Mann-Witney, and Spearman's rank correlation coefficient. Statistical significance was taken at a P value of <0.05 and is denoted in the figures with an asterisk.
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Results |
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Regulation of DLL4 by hypoxia and angiogenic growth factors. Quantitative PCR was used to study DLL4 expression in five endothelial cell lines, including HUVEC, HAEC, HIAEC, HFAEC, and Endo742, cultured at subconfluence in MCDB131 + 20% FCS + ECGS. The expression of DLL4 was highest in HFAEC, with similar expression in HIAEC and HUVEC. DLL4 expression was lowest in HAEC and Endo742 (Fig. 2A). DLL4 was up-regulated by hypoxia in all these cell lines. Cells with high basal expression (HUVEC, HIAEC, and HFAEC) showed a 2- to 3-fold induction whereas those with low basal expression showed a 6- to 7-fold increase in DLL4 expression (Fig. 2B). HIF-1 and HIF-2 in HUVEC were selectively silenced using siRNA (27). Hypoxic up-regulation of the DLL4 expression was preserved in the mock transfection, scrambled control, and HIF-2 groups, but disappeared in the HIF-1 group, indicating that DLL4 is a HIF-1–regulated gene (Fig. 2C).
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The effects of silencing DLL4 using small interfering RNA. Reverse transcription-PCR (RT-PCR) was used to determine which key components of the Notch pathway were expressed by endothelial cells. HUVECs express all four Notch receptors (Notch1-4), four ligands (Jagged1, Jagged2, DLL1, and DLL4), HES1, HEY1, HEY2, and HES7 (weak), but not DLL3, HES5, or HEYL (Fig. 3A). Two different siRNA duplexes specific for DLL4 were used to modulate the expression of DLL4 in HUVECs; DLL4 knockdown was evident up to 4 days posttransfection.
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The effect of DLL4 silencing on endothelial cell proliferation. The proliferation of HUVECs transfected with siRNA duplexes specific for DLL4 (Duplex1 and Duplex2) was significantly inhibited compared with mock transfection and scrambled control (P < 0.001; Fig. 4A). This effect was also shown in Endo742 (data not shown), but not in RT112 (Fig. 4B), a transitional cell carcinoma cell line which does not express DLL4. The inhibition of proliferation was further confirmed using a [3H]thymidine uptake assay (data not shown).
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Analysis of HUVECs stained with propidium iodide showed that DLL4 knockdown led to cell cycle arrest in G0-G1 (Fig. 5A). After serum and growth factor stimulation, 28.3% (±9.3 SE) of cells transfected with scrambled control were in S phase compared with 2.54% (±0.8 SE) for Duplex1 (P < 0.01) and 10.4% (±3.4 SE) for Duplex2 (P < 0.05; Fig. 5B). Down-regulation of DLL4 increased levels of p21 and decreased levels of the phosphorylated retinoblastoma. No change was noted for the p27 protein (Fig. 5C).
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Discussion |
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Although DLL4 was undetectable in adult endothelium in vivo, we found, as have others, that several primary endothelial cells do express it in vitro (29, 33–35). The ratio of DLL4 to CD34 in HUVECs is similar to that found in the highest expressing CC-RCCs. We therefore chose HUVEC as a model to investigate the biological function of DLL4 in tumor angiogenesis. We confirmed that DLL4 is a hypoxia-regulated gene and proved that the hypoxic regulation is dependent on HIF-1 rather than HIF-2.
We showed that the expression of DLL4 in HUVECs is up-regulated by VEGF and bFGF, and their combination leads to a synergistic further increase in expression. Indeed, the combination of VEGF and bFGF can induce DLL4, Notch1, and Notch4 in HUVECs (29). The regulation of DLL4 by VEGF in vitro and the significant correlation between DLL4 and VEGF expression in CC-RCCs imply that VEGF may act upstream of DLL4 in the tumors. It is noted that VEGF can act upstream of DLL4 and Notch4 in the determination of arterial endothelial cell fate in zebra fish (36). bFGF is also produced by renal cancers (37) through a non-HIF pathway. Thus, the combination of VEGF and bFGF may contribute to the high expression of DLL4 within the tumor vasculature.
Using conventional RT-PCR, we showed that HUVECs express all four Notch receptors, four ligands (Jagged1, Jagged2, DLL1, and DLL4), and four bHLH transcription factors [HES1, HEY1, HEY2, and HES7 (weak)], but not DLL3, HES5, and HEYL. Indeed, other groups (29, 35) have shown that HUVECs express Notch1, Notch4, and DLL4, whereas there is also a report indicating that HUVECs express Notch1, 2, 3, and 4 and Jagged1, but not DLL4 (33). The discrepancy for the expression of Notch receptors and ligands in HUVECs may reflect cell heterogeneity or variations between cell batches.
Several groups have attempted to unravel the role of the Notch pathway in endothelial cell biology via the overexpression of Notch signaling components (Jagged1, HES1, and HEY1) or the active Notch intracellular domain of Notch1 and Notch4 (29, 33, 35, 38, 39). Clearly, there are problems of induced high expression, particularly relating to physiologic significance. Therefore, we assessed the biological function of DLL4 by using RNAi to reduce its endogenous expression. Down-regulating DLL4 by 80% led to significant decreases in the expression levels of EphrinB2 and HEY1, indicating a basal role of the DLL4 expression in the regulation of these pathways. HUVECs express several Notch ligands (Jagged1, Jagged2, DLL1, and DLL4); however, regulation of DLL4 alone had substantial effects, suggesting that it is one of the main ligands regulating basal Notch function in endothelial cells. Surprisingly, we have shown that silencing DLL4 in endothelial cells leads to a significant inhibition of proliferation, a result previously found for overexpression studies. The inhibition is due to the arrest of cell cycle in G0-G1, an effect mediated by an increase in levels of p21 and a decrease in the phosphorylation of retinoblastoma. The induction of p21 and resultant decreased phosphorylation of retinoblastoma have previously been shown to induce G0-G1 cell cycle arrest in HUVECs (40–42). However, overexpression of Notch4 or knockout of Notch1 and Notch4 produces similar phenotypes (43, 44). Taken together, this suggests that an optimal range of the Notch signaling is important in the proliferation of endothelial cells.
The effects on apoptosis were reciprocal. We showed that silencing DLL4 induces apoptosis in HUVECs, only following deprivation of serum and growth factors. Overexpression of Notch1, Notch4, or HES1 protects primary endothelial cells against serum starvation–induced apoptosis and inhibits lipopolysaccharide-induced apoptosis (29).
We showed that down-regulation of DLL4 reduces the network formation of HUVECs on matrigel. Up-regulation of Notch1 or HES1 results in stabilization of network formation, whereas inhibition of Notch signaling with a dominant-negative CSL partially inhibits VEGF-driven network formation in HIAECs (29). However, both overexpression and reduction of HEY1 with antisense oligonucleotides blocked endothelial cell network formation in vitro (45). Using the trans-well migration assay and the scratch wound technique, we showed that DLL4 knockdown leads to significant inhibition of VEGF-driven endothelial cell migration. Activated Notch4 inhibits HMEC-1 migration across collagen in response to VEGF and FGF (46), but not through fibrinogen, by increasing the activity and affinity of surface-bound ß1-integrin (47), whereas overexpression of HES1 in endothelial cells decreases migration in response to VEGF or bFGF (45). Thus, again the surprising result is that the effects of down-regulation of DLL4 are similar to the effects of up-regulation of Notch signaling.
In summary, we have quantitatively shown for the first time that the expression of DLL4 is up-regulated in CC-RCCs and that the reduction of basal DLL4 expression has profound effects on multiple endothelial functions important in tumor angiogenesis. Down-regulation of DLL4 produces many phenotypic features displayed by up-regulation of the Notch pathway. Therefore, we propose that an optimal window of the DLL4 expression is essential for tumor angiogenesis. The selective modulation of the DLL4 expression within human tumors may represent a potential novel antiangiogenic therapy.
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Acknowledgments |
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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.
We thank Cassin Kimmel, Richard Poulsom, Mike Dobby, Sarah Pratap, and Roy Bicknell for their help and advice.
Received 4/ 8/05. Revised 5/27/05. Accepted 6/17/05.
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