Introduction

Angiogenesis results from a quantitative, qualitative and temporal balance of cell proliferation, migration and differentiation. Although the mechanism by which this balance is achieved has not been elucidated, some of the signaling enzymes that are required for the angiogenic program have been identified (Matsumoto and Claesson-Welsh, 2001). For instance, phosphoinositide 3 kinase (PI3K) and its downstream effector Akt are essential for many of the cellular responses within the angiogenic program including proliferation, migration and survival (Gerber et al, 1998; Gille et al, 2000; Adini et al, 2003). Similarly, phospholipase C (PLC) is required for vascular endothelial cell growth factor (VEGF)-dependent migration and proliferation of cultured endothelial cells (Takahashi et al, 2001). Furthermore, genetic studies have demonstrated that PLC is essential for angiogenesis (Lawson et al, 2003). PLC-deficient mice die during embryogenesis and display a 'vascular' defect (Liao et al, 2002). Moreover, knocking in a VEGF receptor 2 (VEGFR2) mutant that cannot recruit or activate PLC results in lethality between embryonic days 8.5 and 9.5 and defective hematopoietic development (Sakurai et al, 2005). Although these studies clearly show that PLC is essential for angiogenesis, its precise contribution to the angiogenic program is unknown. Thus, although some of the relevant signaling enzymes have been identified, their contribution to each step of the angiogenic program and their participation in the overall coordination of events remain an open question.

The transition of a stable vessel to one that is capable of entering the angiogenic program is of interest both scientifically and therapeutically. This destabilization step is an early event that is required for all subsequent phases of the angiogenic program and for the regression of vessels (Carmeliet, 2000). At the present time, little is known regarding the molecular nature of this decision or the intracellular signaling events that direct this choice. This information will be invaluable for the treatment of angiogenic diseases (such as solid tumors, diabetic complications and age-related macular degeneration) because current antiangiogenic therapies seem to target immature/unstable vessels (Rasmussen et al, 2001; Shimizu and Oku, 2004). Being able to induce this state will be an important complement to target and potentiate antiangiogenic intervention.

Results and discussion

Preventing activation of PLC enhanced the tube response

One strategy to elucidate the molecular nature of the vessel regression is to identify the relevant signaling enzymes and to determine how they are regulated. To this end, we used an in vitro angiogenesis assay to investigate the role of PI3K and PLC on tube formation and regression. When cultured between two layers of collagen in the presence of VEGF-A, primary bovine retinal endothelial cells (BRECs) formed tubes (Figure 1A and B) (Im et al, 2005). The tubes spontaneously regressed, despite the presence of freshly added VEGF-A (Figure 1A and B). As outlined below, we suspected that PLC promoted regression of tubes. Consequently, we compared vessel formation and regression in BRECs expressing either the wild-type (WT) or mutant VEGFR2 that did not activate PLC (Y1175F) (Takahashi et al, 2001). Somewhat more tubes were consistently observed in the cells expressing the mutant VEGFR2 (Figure 1C). The more striking observation was that the Y1175F tubes did not regress as did the WT tubes (Figure 1C). We repeated these studies with a second primary endothelial cell type (human umbilical vein endothelial cells (HUVECs)) and observed the same phenomenon (Supplementary Figure S1A–C). Furthermore, as an alternative approach to attenuate the contribution of PLC, we used siRNA to reduce the amount of PLC (Figure 1G). Tubes that formed from these cells were resistant to regression, whereas cells expressing a control siRNA (which did not reduce the level of PLC; Figure 1G) regressed normally (Figure 1E). These observations were not unique to the VEGF-driven setting; PLC-dependent regression was also a feature of tubes formed in response to basic fibroblast growth factor (bFGF) (Figure 1F). These findings indicate that the in vitro assay is a model for both tube formation and regression. Furthermore, proangiogenic factors (such as VEGF-A and bFGF) engage a PLC-dependent signaling pathway that promotes tube regression.

PI3K was required for tube formation, whereas PLC promoted tube regression

To investigate the role of PI3K and its relationship with PLC in the context of tube formation and regression, we developed a model system that allowed the conditional activation of each of these signaling enzymes. More specifically, we used platelet-derived growth factor receptor (PDGFR) WT or PDGFR phosphorylation site mutants that do or do not activate PI3K and/or PLC. A comparable set of VEGFR2 or FGFR mutants is not available. BRECs do not express endogenous PDGFRs (Supplementary Figure S5A), although they do express PDGF (Supplementary Figure S2A and B). Stable expression of the WT PDGFR resulted in the spontaneous formation of tubes that were of similar morphology and length to the tubes that formed in parental cells stimulated with VEGF-A (Figure 2A and B). Tube formation was dependent on the extracellular PDGF, as adding a neutralizing PDGF antibody blocked the tube response (Supplementary Figure S2C and D). Supplementing the PDGFR-expressing cells with VEGF-A further enhanced the tube response (Figure 2A and B). Thus, expressing the PDGFR established an autocrine loop that drove spontaneous tube formation.

In certain tumor cell lines, PDGF can stimulate the production of VEGF (Wang et al, 1999). We tested if the tubes that formed in the PDGFR-expressing cells were driven by endogenously produced VEGF. Despite the use of several experimental approaches, we were unable to detect any evidence for such a scenario. Soluble extracellular VEGFR2 blocked VEGF-A-dependent tube formation in parental cells, but had no effect on the tubes that were formed by WT PDGFR-expressing cells (Supplementary Figure S2E). Furthermore, expression of the WT PDGFR did not alter the level of VEGF-A mRNA (Supplementary Figure S2F). We conclude that the PDGF/PDGFR-driven system induced a comparable tube response to that observed when parental cells were stimulated with VEGF-A.

The PDGF/PDGFR-driven system allowed us to evaluate the role of PI3K in tube formation without global inhibition of PI3K activity. A PDGFR mutant that was unable to recruit PI3K (F40/51) failed to induce tubes (Figure 2C). Similarly, the F5 mutant, which lacks bindings sites for PI3K, RasGAP, SHP-2 and PLC, was unable to drive tube formation (Figure 2C). The tube response was restored when we repaired the binding sites for PI3K in the F5 mutant to generate the Y40/51 receptor (Figure 2C). Thus, PI3K was required for tube formation in the PDGF/PDGFR-driven system. Importantly, the same requirement for PI3K was observed with HUVECs stimulated with VEGF-A or bFGF; a PI3K inhibitor blocked tube formation stimulated by either of the proangiogenic factors (Figure 2D). These findings indicated a critical role for PI3K in tube formation, which is consistent with previous reports (Gerber et al, 1998; Hamada et al, 2005).

We routinely noticed that tube formation with Y40/51 cells was better than WT cells (Figure 2C). This suggested that one or more of the signaling enzymes recruited by the WT receptor (RasGAP, SHP-2 or PLC) were suppressing the response. Given the known involvement of PLC in angiogenesis (Takahashi et al, 2001; Liao et al, 2002; Lawson et al, 2003) and our findings in the VEGF-A- and bFGF-driven systems (Figure 1C, E and F and Supplementary Figure S1B), we focused on PLC. Tube formation in cells expressing a receptor that did not activate PLC (F1021) was better than in cells expressing the WT receptor (Figure 2E). The same trend was observed in HUVECs stimulated with either VEGF or bFGF; reducing the level of PLC with siRNA boosted tube formation (Figure 2F). These studies revealed that PI3K promoted tube formation, and that PLC negatively impacted one or more steps in the overall tube response.

The negative impact of PLC on the overall tube response could be from blocking tube formation and/or promoting regression of tubes after they have formed. To investigate this issue, we examined both tube formation and regression using PDGFR mutants that activated either or both PI3K and PLC. Within the first day of the experiment, tubes formed comparably for both the Y40/51 and Y40/51/21 cells (Figure 2G), indicating that PLC did not prevent the formation of tubes. Instead, PLC appeared to promote tube regression because at the latter time points the Y40/51/21 tubes regressed, whereas the Y40/51 tubes persisted (Figure 2G). Similarly, cells expressing PDGFR mutants that recruited PI3K in combination with RasGAP or SHP-2 (instead of PLC) formed stable tubes (Supplementary Figure S2G). These experiments indicate that PLC promoted regression.

Taken together, the results in Figure 2 indicate that PI3K was essential for tube formation, whereas PLC reduced the overall tube response by inducing tube regression.

Activation of PLC reduced the output of PI3K

Our findings in Figure 2 suggested an antagonistic relationship between PI3K and PLC. These two enzymes require the same substrate (phosphatidylinositol-4,5-bisphosphate (PtdIns-4,5-P₂)) raising the possibility that the antagonism resulted from a competition for substrate. More specifically, we speculated that activation of PLC reduced the level of PtdIns-4,5-P₂ available for PI3K and thereby attenuated the output of the PI3K pathway.

A prediction of the substrate competition hypothesis is that phosphoAkt (a downstream target of PI3K) would be reduced in situations where PLC is activated. This is what we observed using several experimental settings. In PDGF- or VEGF-stimulated monolayers, phosphoAkt was lower in cells expressing receptors that activated PLC as compared with receptors that did not (Figure 3A and Supplementary Figure S3A). A similar phenomenon was observed when the cells were organized into tubes; the extent of phosphoAkt was greater when the receptor (PDGFR or VEGFR2) failed to activate PLC or when the level of PLC was reduced with siRNA (Figure 3B and Supplementary Figure S3D–F). For the latter time points (Figure 3B and Supplementary Figure S3B), the tubes were harvested before regression, and so the decline in phosphoAkt was not the result of regression. Furthermore, other signaling systems (PLC tyrosine phosphorylation) did not fall at the latter time points, arguing against a global decline in signaling (Figure 3C and Supplementary Figure S3C). Thus, all three experimental approaches indicate that PLC antagonized PI3K/Akt signaling.

PLC is linked to Erk activation in VEGF-stimulated endothelial cells (Takahashi et al, 2001). Yet in our hands, Erk activation was not significantly impaired when we interfered with PLC (Supplementary Figure S3G–I). This may be because neither of the experimental systems completely eliminated PLC activation, and thus the remaining PLC activity may have been sufficient to fully activate Erk.

A second prediction of the substrate competition hypothesis is that boosting the cellular level of PtdIns-4,5-P₂ would restore the output of the PI3K pathway. Indeed, adding synthetic PtdIns-4,5-P₂ to Y40/51/21 tubes increased phosphoAkt to the day 1 level (Figure 3D). The finding that this response was blocked by a PI3K inhibitor (LY294002) (Figure 3D) further supported the idea that PLC created a shortage of PtdIns-4,5-P₂. Adding the PI3K lipid product (PtdIns-3,4,5-P₃) also prevented the fall in phosphoAkt, but this response was not sensitive to PI3K inhibitors (Figure 3D), as it was downstream of PI3K. The results in Figures 3 and Supplementary Figure S3 support the hypothesis that PLC antagonized the PI3K pathway at the level of lipid substrate.

Synthetic PI3K lipid substrates prevented tube regression

We also tested if increasing the cellular level of PtdIns-4,5-P₂ prevented regression of tubes. In both the PDGF/PDGFR and VEGF/VEGFR systems, PtdIns-4,5-P₂ stabilized tubes and this event was sensitive to inhibition of PI3K activity (Figure 4). PtdIns-3,4,5-P₃ also stabilized tubes, but in this case the PI3K inhibitor had no effect. Time-lapse photography of the tubes indicated that the synthetic lipids did not induce new tube formation, but rather prevented existing tubes from disappearing (unpublished observation). We concluded that, like the decline in phosphoAkt, the PLC-mediated regression of tubes resulted from an attenuation of the PI3K/Akt output.

PI3K/Akt promoted survival of vessels (Gerber et al, 1998; Adini et al, 2003), which prompted us to consider whether tube regression was the result of apoptosis. Three different measures of apoptosis (cell number, caspase 3 cleavage and TUNEL staining) indicated that apoptosis was occurring (Supplementary Figure S4). However, the tubes regressed before they underwent apoptosis. When we chemically induced apoptosis in stable tubes using a high concentration of a PI3K inhibitor, it resulted in apoptotic cells geographically arranged in the tube structures. Whereas, when the tubes regressed in our system, the cells within the tubes retracted into aggregates, and then underwent apoptosis (Supplementary Figure S4C). As the tubes collapsed into aggregates, there was an expected change in the actin (phalloidin staining; Supplementary Figure S4D), which may be casually related to apoptosis. Thus, although the cells in regressing tubes did eventually undergo apoptosis, it appeared that apoptosis was not the cause for regression. Others have also reported that tubes/vessels first regress, and then apoptose (Bayless and Davis, 2004; Saunders et al, 2005).

Partial inhibition of PI3K induced tube regression without affecting tube formation

The competition hypothesis provided an explanation for why tubes regressed, but left open the question of why they were capable of forming initially. One possibility was that the antagonism (i.e. PLC activation) was engaged only after tubes formed. However, Figure 3C and Supplementary Figure S3C show comparable tyrosine phosphorylation of PLC as tubes formed and regressed. Thus, the degree of PLC-dependent antagonism appeared to be constant. A second possibility was that more PI3K signaling was necessary for stabilization of tubes as opposed to their formation. Thus, PLC-dependent attenuation of PI3K output was insufficient to prevent the tubes from forming. This idea is consistent with previous findings that PI3K inhibitors promoted tube regression without affecting their formation using endothelial cells of the adrenal cortex (Qi et al, 1999). To test this idea, we compared the effect of sub-maximal doses of LY294002 on vessel formation and stability. Partially blocking PI3K (using 2 or 5 M of LY294002) had no effect on tube formation, yet it was sufficient to induce tube regression (Figure 5). Higher doses of LY294002 inhibited formation of tubes in BRECs and HUVECs (Figures 5 and 2D), which was consistent with the PDGFR mutant data showing that PI3K was necessary for tube formation (Figure 2C). These observations suggested that tube stability was more dependent on PI3K signaling than the tube formation phase of the angiogenic program.

The data presented support the idea that PLC promoted vessel regression by antagonizing PI3K. However, they did not rule out other possibilities. For instance, PLC may promote the synthesis/secretion of antiangiogenic factors, whose input is masked by flooding the system with synthetic lipids. Although our studies leave open this (and other) possibilities, they do strongly suggest that the amount of PtdIns-4,5-P₂ is limiting, and support our overall conclusion that the angiogenic program can be regulated at the level of a membrane lipid.

Our finding that PLC controls tube formation by reducing the cellular level of PtdIns-4,5-P₂ further increases our appreciation of how PtdIns-4,5-P₂ contributes to cell signaling. It is not only a depot for the protein kinase C family activator (diacylglycerol) and intracellular calcium booster (inositol triphosphate) (Irvine, 2000), it can also inhibit tyrosine kinases (c-abl) (Plattner et al, 2003) and thereby modify a variety of cellular responses. Finally, the data presented herein indicate that PtdIns-4,5-P₂ can act as an interface between PI3K and PLC to coordinate the various phases of the angiogenic program.

Materials and methods

Antibodies and reagents

Anti-mouse and anti-rabbit antibodies conjugated to horseradish peroxidase were obtained from Amersham Biosciences (Piscataway, NJ). Rabbit polyclonal anti-phospho-Akt antibody and anti-Akt antibody were obtained from Cell Signaling Technology (Beverly, MA). Mouse monoclonal anti-phosphotyrosine antibodies PY20 and 4G10 were purchased from Transduction Labs (Lexington, KY) and Upstate Biotechnology Inc. (Lake Placid, NY), respectively. The RasGAP, PDGFR, PLC and VEGFR2 antibodies were crude polyclonal rabbit antisera that were previously described (Valius et al, 1993; Rahimi and Kazlauskas, 1999). LY294002 was purchased from Calbiochem (San Diego, CA). The WT VEGFR2 and Y1175F VEGFR2 cDNAs were kindly provided by Dr Lena Claesson-Welsh (Uppsala University, Uppsala, Sweden). Recombinant VEGF-A was purchased from Upstate Biotechnology Inc. PDGF BB was purchased from R&D system (Minneapolis, MN). All other chemicals and reagents were obtained from Sigma (St Louis, MO) unless otherwise indicated.

Cell culture

BRECs were isolated from bovine eyes as described previously (Gitlin and D'Amore, 1983; Im et al, 2005). BRECs were maintained in EBM (Clonetics, Walkersville, MD) supplemented with 10% horse serum (Clonetics), 80 U/ml penicillin/streptomycin C (Irvine Scientific, Santa Ana, CA), and 12 g/ml bovine brain extract (Clonetics). The cells were plated on plastic coated with 50 g/ml bovine fibronectin and incubated at 37°C in 5% CO₂. For all experiments, cells were used between passages 7 and 10. HUVECs were purchased from Clonetics and maintained in EGM-2 (Clonetics) with low serum growth factor supplement (Clonetics). For all experiments, HUVECs were used between passages 5 and 7.

Tube formation assay

Tube formation assay was performed as previously described (Im et al, 2005). The average tube length was routinely 15–30 mm in either BRECs or HUVECs exposed to VEGF-A, or bFGF, or PDGFR-expressing BRECs responding to endogenous PDGF.

Transfection of siRNA-PLC oligonucleotides

siRNA oligonucleotides that target PLC and non-targeting siRNA pool were purchased from Dharmacon (Lafayette, CO) and resuspended according to the manufacturer's instructions. For transfection, 1 10⁵ HUVECs were plated into each well of a six-well dish and incubated for 16–18 h in culture medium. A 100 nM portion each of siRNA-PLC and siRNA-control oligonucleotides was mixed with TransPass™ R2 transfection reagent (New England BioLabs, Beverly, MA) 20 min before transfection. Cells were washed once with DMEM (GIBCO BRL, Gaithersburg, MD) and then the transfection reagent mixture was added. After a 4 h incubation, 2 ml of culture medium was added and the cultures were incubated overnight. Cells were then incubated for 48 h in freshly added culture medium.

Stable expression of PDGFR mutants

The WT, F40/51, F1021, Y40/51, Y1021, Y40/51/21 and F5 PDGFR constructs were previously constructed and characterized (Valius and Kazlauskas, 1993). Briefly, the F5 receptor has tyrosines 740, 751, 1009 and 1021 mutated to phenylalanine. The Y40/51 receptor is the same as F5, except that the mutations at 740 and 751 have been repaired. Similarly, the Y1021 receptor has tyrosine in place of phenylalanine at position 1021, whereas all the other mutations are as in F5. The F5 PDGFR does not efficiently associated with PLC, RasGAP, PI3K or SHP-2 and fails to activate either PI3K or PLC. The Y40/51 and Y1021 PDGFRs recruit PI3K and PLC, respectively. The PDGFR cDNAs were subcloned into the retroviral vector pLXSN. The PLSXN empty vector and PDGFR mutants/pLXSN constructs were transfected into 293GPG cells. The supernatant was collected for 5 days, concentrated (25 000 g, 90 min, 4°C) and used as described previously (Ory et al, 1996). Cells were infected and selected on the basis of proliferation in the presence of G418 (1 mg/ml).

Western blot analysis and immunoprecipitation

For Western blot analysis of VEGFR2, 2 10⁶ BRECs or HUVECs were plated into a 10 cm tissue culture plate and incubated for at least 18 h in culture medium. Total cell lysates were prepared by adding lysis buffer containing 20 mM Tris (pH 8.0), 150 mM NaCl, 1 mM dithiothreitol, 1% deoxycholic acid, 0.5% sodium dodecyl sulfate (SDS), 1% Nonidet P-40 and protease inhibitors (2 g/ml aprotinin, 5 g/ml leupeptin, 10 g/ml phenyl methyl sulfonyl fluoride and 10 mM sodium fluoride) and incubating for 1 h on ice. After centrifugation, the supernatants were collected and the protein concentration was determined. Proteins (10–30 g) were separated on 10% SDS–polyacrylamide gels, and Western blot analysis was performed as described previously (Im et al, 2005).

To reprobe a blot, the blot was first stripped by incubating for 30 min at 60°C in a buffer containing 6.25 mM Tris–HCl, pH 6.8, 2% SDS and 100 mM -mercaptoethanol and then reprobed with the desired primary antibody.

For Western blot analysis of phospho-Akt and Akt, the PDGFR-expressing cells or VEGFR2 WT/Y1175F-expressing cells or siRNA PLC/control siRNA-transfected cells were plated in a collagen sandwich gel and incubated for the desired time period. The cells were recovered following a collagenase treatment (collagenase type I-S from Sigma, 281 U/well for 20 min at 37°C), which dissolved the collagen gel. The cells were rinsed three times with ice-cold PBS, total cell lysates were made and then subjected to Western blot analysis as described above.

To analyze tyrosine phosphorylation of PLC in the tubes, cells were recovered from tubes and lysed as described above. The lysate was precleared with non-immune antibodies coupled to protein A/G agarose (Santa Cruz Biotechnology, Santa Cruz, CA), and then PLC was immunoprecipitated. Immunocomplexes were collected on protein A/G plus agarose and washed three times with the lysis buffer described above. The immunoprecipitated proteins were subjected to an anti-phosphotyrosine Western blot as described above.

Synthetic lipids rescue assay

The delivery of synthetic lipids to the cells was performed as previously described (Weiner et al, 2002). Briefly, parental or PDGFR-expressing BRECs were plated in a collagen sandwich gel and incubated for 12 h before adding lipids. Di-C16 synthetic phospholipids PtdIns-4,5-P₂ and PtdIns-3,4,5-P₃ (Echelon, Salt Lake City, UT) were freshly prepared at 25 M in 150 mM sodium chloride, 4 mM potassium chloride and 20 mM HEPES at pH 7.2, and resuspended by vigorous vortexing. For histone–phospholipid complexes, 25 M phospholipids was mixed with 100 M freshly prepared histone (Echelon), vortexed vigorously and incubated for 5 min at room temperature. Histone–phospholipid complexes were diluted 1:10 with modified Hanks buffered saline solution immediately before addition to the media on top of the collagen gel. These media were replaced every 12 h until the end of the experiment.

Statistics

The Student's t-test was used to assess statistical significance.

Supplementary data

Supplementary data are available at The EMBO Journal Online.

Acknowledgements

We thank Dr Lena Claesson-Welsh for kindly providing the WT and Y1175F VEGFR2 cDNAs. We also thank Drs Laura E Benjamin, Chris Carpenter, Lena Claesson-Welsh, Giulio Romeo, Rita N Barcia, Hetian Lei and Sang Hoon Rhee for critically reading the manuscript. This study was supported by a Pediatric Ophthalmology Research Grant of the Knights Templar Eye Foundation Inc. (to EI), and by an NIH grant EY016385 (to AK).

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