October 2009
Volume 50, Issue 10
Free
Biochemistry and Molecular Biology  |   October 2009
FAK and p38-MAP Kinase-Dependent Activation of Apoptosis and Caspase-3 in Retinal Endothelial Cells by α1(IV)NC1
Author Affiliations
  • Chandra S. Boosani
    From the Cell Signaling and Tumor Angiogenesis Laboratory and the
  • Narasimharao Nalabothula
    From the Cell Signaling and Tumor Angiogenesis Laboratory and the
  • Veerendra Munugalavadla
    Department of Pediatrics, Indiana University School of Medicine, Indianapolis, Indiana;
  • Dominic Cosgrove
    Gene Expression Laboratory, Department of Genetics, Boys Town National Research Hospital, Omaha, Nebraska;
  • Venkateshwar G. Keshamoun
    Division of Pulmonary and Critical Care Medicine, Department of Internal Medicine, University of Michigan Medical Center, Ann Arbor, Michigan;
  • Nader Sheibani
    Department of Ophthalmology and Vision Science, University of Wisconsin School of Medicine and Public Health, Madison, Wisconsin;
  • Akulapalli Sudhakar
    From the Cell Signaling and Tumor Angiogenesis Laboratory and the
    Department of Biomedical Sciences, Creighton University School of Medicine, Omaha, Nebraska; and
    Department of Biochemistry and Molecular Biology, University of Nebraska Medical Center, Omaha, Nebraska.
Investigative Ophthalmology & Visual Science October 2009, Vol.50, 4567-4575. doi:10.1167/iovs.09-3473
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      Chandra S. Boosani, Narasimharao Nalabothula, Veerendra Munugalavadla, Dominic Cosgrove, Venkateshwar G. Keshamoun, Nader Sheibani, Akulapalli Sudhakar; FAK and p38-MAP Kinase-Dependent Activation of Apoptosis and Caspase-3 in Retinal Endothelial Cells by α1(IV)NC1. Invest. Ophthalmol. Vis. Sci. 2009;50(10):4567-4575. doi: 10.1167/iovs.09-3473.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. To determine the impact of the antiangiogenic factor α1(IV)NC1 on vascular endothelial growth factor–mediated proangiogenic activity in mouse retinal endothelial cells (MRECs).

methods. Primary culture of MRECs was established as previously described and was used to determine the effects of α1(IV)NC1 on the proangiogenic activity of VEGF. Cell proliferation was evaluated using [3H]-thymidine incorporation and 3,(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide colorimetric assays. Cell migration was determined using modified Boyden chamber and scratch wound assays and tube formation was assessed on basement membrane matrix (BMM). Intracellular signaling events Bcl-2/Bcl-xL and caspase-3/poly (ADP-ribose) polymerase (PARP) activities were evaluated in cells stimulated with VEGF and plated on type IV collagen-coated dishes. Apoptosis was assessed by measuring caspase activity and by performing quantitative fluorescence analysis using fluorescence-activated cell sorting assay. Subcutaneously injected VEGF induced in vivo neovascularization was studied with the BMM plug assay.

results. VEGF-induced subconfluent MREC proliferation, migration, and tube formation were significantly inhibited by α1(IV)NC1 at 1 μM (P < 0.001). α1(IV)NC1 induced MREC apoptosis is mediated by inhibition of Bcl-2 and Bcl-xL expression and activation of caspase-3/PARP through FAK/p38-MAPK signaling. In addition, α1(IV)NC1 dose dependently inhibited VEGF-mediated neovascularization in vivo.

conclusions. α1(IV)NC1 inhibited VEGF-mediated angiogenesis by promoting apoptosis and caspase-3/PARP activation and by negatively impacting FAK/p38-MAPK phosphorylation, Bcl-2, and Bcl-xL expression leading to MREC death. The endothelial-specific inhibitory actions of recombinant α1(IV)NC1 may be of benefit in the treatment of a variety of eye diseases with a neovascular component.

Angiogenesis, the formation of new blood vessels from preexisting capillaries, is a tightly regulated process and normally does not occur, except during developmental and wound repair processes. 1 This strict regulation is manifested by a balanced production of proangiogenic and antiangiogenic factors that keep angiogenesis in check. 1 However, this balance between proangiogenic and antiangiogenic factors abrogates under many pathologic conditions, including cancer, diabetes, age-related macular degeneration, and retinopathy of prematurity, resulting in the growth of abnormal new blood vessels. 2 3 4  
Vascular endothelial growth factor (VEGF) is a major proangiogenic factor that promotes the growth of new blood vessels under ischemic conditions. VEGF stimulates endothelial cell proliferation, migration, and survival. When retinal pigment epithelial cells begin to wither because of lack of nutrition (ischemia), VEGF goes into action to create neovascularization, and it acts as a restorative function in other parts of the body. In the retina, these vessels do not form properly and leak, ultimately resulting in retinal detachment and loss of vision. Recently, several antiangiogenic compounds have been shown to inhibit neovascularization and prevent bleeding into the vitreous by inhibiting VEGF binding to its receptors on endothelial cells. 5 6 Therefore, there is significant interest in the development and identification of molecules that can inhibit the growth of new blood vessels. 
Vascular basement membrane (VBM) constitutes an important component of blood vessels. 7 Remodeling of VBM can provide crucial proangiogenic and antiangiogenic molecules to control the formation of new capillaries. 8 9 10 Type IV collagen is a major component of VBM and plays a crucial role in the regulation of angiogenesis. 7 Proteolytic degradation of type IV collagen in the VBM generates antiangiogenic molecules. 11 One such antiangiogenic molecule, derived from the α1 chain of type IV collagen noncollagenous (NC1) domain, α1(IV)NC1 (arresten), has been tested in mice against a variety of cancers. 12 13 However, the molecular and cellular mechanisms responsible for the inhibition of angiogenesis require further delineation. In vitro and in vivo studies have demonstrated that α1(IV)NC1 can directly affect endothelial cell migration and impact their proliferation and sprouting. 13 However, the effects of α1(IV)NC1 on retinal endothelial cell function and vascularization have not been previously studied. 
In the present study, we demonstrate that α1(IV)NC1 is a potent inhibitor of mouse retinal endothelial cell (MREC) proliferation, migration, and tube formation in vitro and angiogenesis in vivo. We demonstrate that α1(IV)NC1 promotes apoptosis through the activation of caspase-3 and PARP cleavage, presumably by inhibiting FAK/p38-mitogen-activated protein kinase (MAPK)/Bcl-2 and Bcl-xL signaling cascades in MREC without affecting mouse retinal pigment epithelial (MRPE) cells. These findings contribute significantly to our understanding of the apoptotic signaling mechanism and therapeutic potential of the α1(IV)NC1 molecule in retinal diseases with a neovascular component. 
Methods
Preparation and Culture of Primary Mouse Retinal Endothelial and Retinal Pigment Epithelial Cells
All experiments were conducted in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. Primary MRECs were prepared and maintained as described previously. 14 MRECs were maintained in 40% HAM F-12, 40% DME-low glucose, 20% FCS supplemented with heparin (50 mg/L), endothelial mitogen (50 mg/L), l-glutamine (2 mM), penicillin/streptomycin (100 U/mL each), Na pyruvate (2.5 mM), NEAA (1×), and 5 μg/L murine IFN-γ and were cultured on 0.8% gelatin-coated plates at 33°C with 5% CO2. MRPE cells were maintained in DMEM containing 10% FCS and 100 U/mL antibiotic and antimycotic solutions at 37°C with 5% CO2
Proliferation Assay
A suspension of 40 × 103 MRECs/well was used in the proliferation assay. MREC medium (500 μL/well) was added to 24-well plates precoated with 10 μg/mL type IV collagen. After 24 hours, that medium was replaced with MREC medium containing 10 ng/mL VEGF, and different concentrations of baculovirus expressed recombinant α1(IV)NC1 or α3(IV)NC1 (tumstatin) (0.5 and 1.0 μM). After 24 hours, 1 μL [3H]-thymidine was added into each well. After 24 hours, [3H]-thymidine incorporation was measured with a scintillation counter. 13  
Cell Viability Assay
The viability of MRECs and MRPE cells was assessed by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide assay (MTT) according to the manufacturer’s protocols. Overnight serum-starved MRECs or MRPE cells (7.0 × 103/well) were plated on a 96-well plate and stimulated with VEGF (10 ng/mL)–containing medium. The next day, different concentrations of α1(IV)NC1 (0.25–1.0 μM) were added and incubated for 24 hours. Apoptosis was monitored by trypan blue exclusion using the cell death detection ELISA kit. 15  
Migration Assay
MRECs (1.0 × 104/well) were seeded in no-serum medium with and without recombinant α1(IV)NC1. Medium containing 10 ng/mL VEGF was placed in the bottom wells of the Boyden chamber and incubated for 24 hours at 37°C with 5% CO2. The numbers of MRECs that migrated and attached to the bottom side of the membrane were counted. 13 16  
Tube Formation Assay
Briefly, approximately 250 μL basement membrane matrix (BMM; Matrigel; BD Biosciences, San Jose, CA) was added to each well of a 24-well plate and allowed to harden for 30 minutes at 37°C. A suspension of 50 × 103 MRECs in medium without antibiotic was plated on top of the BMM-coated wells. The MRECs were stimulated with VEGF for tube stabilization and incubated with and without α1(IV)NC1 for 24 hours at 37°C and viewed under a microscope (CK2; Olympus, Tokyo, Japan). 17 18  
Scratch Wound Assays
Briefly, MREC wound assays were conducted in type IV collagen-coated six-well tissue culture plates. MRECs were cultured to subconfluence, and wounds were created at the center of each plate by scraping the monolayer with a sterile pipette tip. 19 Medium containing 1 μM α1(IV)NC1 was added only to experimental wells, and cells migrating to the wound area were monitored microscopically for 36 hours. Photographs were taken at different times, and migrated cells were counted in control and in α1(IV)NC1-treated wells. 
Caspase-3, -8, and -9 Activity Assays
MRECs and MRPE cells were grown to subconfluence; cells detached and were plated at 0.5 × 106 cells/well in a six-well plate (precoated with type IV collagen 10 μg/mL) in 10% FCS-supplemented medium overnight. The following day, serum-containing medium was replaced with serum-free medium and incubated overnight at 37°C. Cells were then stimulated with VEGF (10 ng/mL) in MREC medium containing α1(IV)NC1 (1 μM) and were incubated for 24 hours. TNF-α (80 ng/mL) was used as a positive control for caspase-3–dependent apoptosis, whereas staurosporine (50 nM) was used as a positive control for mitochondrial caspase-dependent apoptosis. A specific inhibitor of caspase-3, DEVD-fmk (20 μM), was used to confirm the specificity of the assay. Similarly, caspase-8 and -9 inhibitors z-IEPD-fmk and z-LEHD-fmk (20 μM) were used. After 24 hours, the floating cells were collected, combined with the adherent cells, and washed three times with sterile PBS. Equal numbers of cells in each condition were resuspended and mixed in appropriate peptide substrate reaction buffers containing 100 mM HEPES, pH 7.0, (for caspase-3 and -8) and 100 mM MES, pH 6.5, (for caspase-9) in 30 μL PBS and were transferred to a 96-well plate for fluorometric analysis in a microplate reader. Caspase-3, -8 and -9 activities were determined fluorometrically by cleavage of cellular substrate DEVD-AMC, IEPD-AMC, or LEHD-AFC, as described. 20  
FACS Analysis of MREC Incubated with α1(IV)NC1
Apoptotic and viable MRECs were measured by quantitative fluorescence analysis performed using fluorescence-activated cell sorting (FACS). Briefly, MRECs were incubated with and without α1(IV)NC1 (1 μM) for 15 minutes at room temperature and were plated onto type IV collagen-coated 10-cm tissue culture plates containing VEGF (10 ng/mL). The culture plates were incubated at 37°C with 5% CO2 for 24 hours and were used for FACS analysis. Floating MRECs were collected, combined with adherent cells, removed by trypsinization (0.05% [wt/vol] trypsin in 0.02% [wt/vol] EDTA), incubated for 5 minutes at 37°C, and washed twice with PBS, and the cell pellet was resuspended in 1× binding buffer (10 mM HEPES/NaOH [pH 7.4], 140 mM NaCl, 2.5 mM CaCl2). In each condition, MRECs were suspended in binding buffer containing fluorescein isothiocyanate (FITC)–conjugated annexin-V and propidium iodide (PI) and were incubated for 20 minutes at room temperature in the dark. Additional binding buffer was added, and 10,000 events were acquired and analyzed by FACS. 21  
Cell Signaling Experiments
For FAK and p38-MAPK signaling experiments, approximately 1.0 × 106 serum-starved MRECs or MRPE cells were preincubated for 10 minutes with α1(IV)NC1 (1 μM), followed by VEGF (10 ng/mL) stimulation, and were seeded into 10-cm2 dishes coated with type IV collagen (10 mg/mL) for different time intervals. Cells were lysed in RIPA lysis buffer, and extracts were analyzed by SDS-PAGE and immunoblotting with antibodies specific to phosphorylated and total FAK or p38-MAPK. 13 18 For measuring caspase-3 activation, MRECs were incubated with α1(IV)NC1 for 6 to 24 hours at 37°C. Floating and adherent cells were pooled and lysed, and cytosolic extracts were immunoblotted. Cleaved activated caspase-3 was identified by specific monoclonal antibodies and with the use of an ECL detection kit. 
Effect of α1(IV)NC1 on p38-MAPK–Dependent Bcl-2/Bcl-xL Regulation and Caspase-3/PARP Activation
Approximately 1 × 106 serum-starved MRECs were collected and suspended in serum-free medium containing VEGF (10 ng/mL). These cells were pretreated with α1(IV)NC1 (1 μM) or p38-MAPK specific inhibitor SB203580 (20 μM) or TNF-α (80 ng/mL) for approximately 10 minutes and were seeded on type IV collagen precoated 10-cm2 dishes for 24 hours. Plates were washed once with cold PBS, and cells were lysed on ice in RIPA lysis buffer and centrifuged at 4°C for 30 minutes at 13000 rpm. Approximately 30 μg cytosolic extract/lane was separated using 10% SDS-PAGE, followed by immunoblotting with anti–Bcl-xL, Bcl-2, caspase-3, and PARP antibodies. Immunoreactivity of Bcl-xL, Bcl-2 proteins, cleaved caspase-3, and PARP were visualized with the ECL detection kit. 
BMM Plug Assay
Fifteen male 12-week-old 129/Sv mice were used for this study. For each condition, three mice or six BMMs were used. All animal studies were reviewed and approved by the animal care and use committee of Boys Town National Research Hospital. BMM was injected subcutaneously into the right and left sides of mice, lateral to the abdominal midline. Control groups received only heparin in the BMM, whereas positive control groups received heparin and VEGF (100 ng/mL) with and without type IV collagen (10 μg/mL). The experimental group received two different doses of α1(IV)NC1 (0.5 and 1 μM) along with heparin and VEGF in the BMM. 22 After 10 days, mice were killed, and half the BMM plugs were fixed in 4% paraformaldehyde, sectioned, and stained with hematoxylin and eosin. The numbers of blood vessels from 12 high-power fields were counted. The other half of the BMM plugs were dispersed in sterile PBS and incubated at 4°C overnight. Hemoglobin levels were determined with Drabkin reagent. 18  
Statistical Analysis
Each experiment was performed three times and values are expressed as mean ± SEM. Statistical analysis based on the Student’s t-test (unilateral and unpaired) was scored to identify significant differences in multiple comparisons. P < 0.05 was considered statistically significant. 
Results
Distinct Antiangiogenic Activities of α1(IV)NC1 on MRECs
α1(IV)NC1 was discovered as an antiangiogenic molecule with significant antitumor activities. 13 α1(IV)NC1 is liberated from the noncollagenous (NC1) domain of the α1 chain of type IV collagen by matrix metalloproteinase-9 (MMP-9). 23 The present studies were aimed at achieving an understanding the molecular mechanisms underlying angiogenesis inhibition by α1(IV)NC1 and its implications in the treatment of cancer and were conducted in an attempt to determine the antiangiogenic actions of α1(IV)NC1 in MRECs. 
A series of angiogenesis experiments was conducted to determine the antiangiogenic activity of α1(IV)NC1 using MRECs and MRPE cells. VEGF is a major proangiogenic factor responsible for most ocular angiogenesis driven by ischemia. We first determined VEGF-stimulated antiangiogenic activity of α1(IV)NC1 by measuring MREC proliferation. We used [3H]-thymidine incorporation to examine the antiproliferative effect of α1(IV)NC1. VEGF-stimulated proliferation of MRECs was significantly inhibited by α1(IV)NC1 in a concentration-dependent manner compared with α3(IV)NC1. The graph summarizes relative [3H]-thymidine incorporation in MRECs incubated with α1(IV)NC1 and α3(IV)NC1 (α1(IV)NC1 70.49% vs. α3(IV)NC1 50.49% at 1 μm; Fig. 1A ). We also confirmed that MREC proliferation was inhibited dose dependently by α1(IV)NC1 with the use of methylene blue staining (Fig. 1B) . Such proliferation inhibition ([3H]-thymidine incorporation or methylene blue staining) was not observed in MRPE cells incubated with α1(IV)NC1 (data not shown). Further, we tested antiangiogenic activity at different doses of α1(IV)NC1 (0, 0.25, 0.5, 0.75, and 1.0 μM) by MTT cell viability assay after VEGF stimulation in MRECs. The results showed that MREC proliferation was increased significantly by VEGF stimulation, which was inhibited by α1(IV)NC1 dose dependently after 24 hours (Fig. 1C) . Interestingly, α1(IV)NC1 had no significant effect on MRPE cell proliferation in the presence of VEGF, indicating the endothelial cell-specific action of α1(IV)NC1 (Fig. 1D)
The migration of endothelial cells is fundamentally important during angiogenesis. 13 24 MREC migration across a collagen type IV–coated membrane toward VEGF in a Boyden chamber was significantly inhibited by α1(IV)NC1 in a dose-dependent manner (0, 0.25, 0.5, and 1.0 μM; Figs. 2A 2B ). We further confirmed the antiangiogenic activity of α1(IV)NC1 by a functional assay of VEGF-mediated, stabilized MREC tube formation. 17 Tube formation on BMM is associated with endothelial cell migration and survival. 25 Incubation of MREC with α1(IV)NC1 inhibited VEGF-stabilized tube formation on BMM in a dose-dependent manner (Fig. 3A , α1(IV)NC1). Our previous report states that the antiangiogenic activity of α1(IV)NC1 is mediated through α1β1 integrin. 13 Interestingly, we observed that MRECs, when incubated with α1(IV)NC1 (1.0 μM), becomes rounded (apoptosis like) and detaches from the BMM, forming clumps of cells (Fig. 3A , lower right corner). This is perhaps caused by activation of the apoptosis in MRECs incubated with α1(IV)NC1. The number of tubes formed after 36 hours in the presence of different doses of α1(IV)NC1 from three independent experiments is shown in Figure 3B
The decrease in the proliferation of MRECs incubated with α1(IV)NC1 was associated with a significant reduction in the migration of these cells. Consistent with the reduced proliferation and migration of MRECs incubated with α1(IV)NC1 on type IV collagen, we also observed the attenuation of MREC migration in scratch wound assays (Fig. 4A , α1(IV)NC1). In addition, we observed apoptotic MRECs during incubation with α1(IV)NC1. Prolonged incubation of MRECs with α1(IV)NC1 resulted in their rounding and detachment from the matrix, indicating the onset of apoptotic events. Moreover, the number of adherent MRECs decreased, and floating cells increased compared with control cells, suggesting that cell detachment was caused by the presence of α1(IV)NC1. The average number of migrated MRECs into the scratch wound at different times with and without α1(IV)NC1 treatment in three independent experiments is shown in Figure 4B
Impairment of In Vitro Apoptotic Activity of α1(IV)NC1 by Caspase Inhibitors
In tracing the signaling mechanisms involved in the rapid impairment of cell proliferation that precedes the loss of MREC viability and the irreversible commitment to cell death on incubation with α1(IV)NC1, we identified that α1(IV)NC1 induces apoptosis in MRECs (Fig. 5A) . To study whether caspase-3 could be activated by α1(IV)NC1, we incubated MRECs with α1(IV)NC1 and observed the activation of caspase-3 (Fig. 5A) . Caspase-3 is a pivotal molecule mediating cellular apoptosis. 26 If this activation of caspase-3 by α1(IV)NC1 is necessary for cellular apoptosis, a caspase-3–specific inhibitor should abolish α1(IV)NC1-induced apoptosis in MRECs. The incubation of MRECs with z-DEVD (a specific caspase-3 inhibitor) showed complete suppression α1(IV)NC1-induced cellular apoptosis and inhibition of caspase-3 activity (Fig. 5A , α1(IV)NC1+DEVD). Similar apoptotic caspase-3 activation was not observed in MRPE cells incubated with α1(IV)NC1 (Fig. 5B , α1(IV)NC1). These results suggest that the proapoptotic action of α1(IV)NC1 through the activation of caspase-3 is specific to MRECs. 
We further evaluated whether α1(IV)NC1 induces the activation of upstream caspases such as caspase-8 and -9. We treated MRECs with α1(IV)NC1 and observed the activation of caspase-8 and -9 (Figs. 5C 5D) . If this activation of caspase-8 and -9 by α1(IV)NC1 plays a role in cellular apoptosis, specific inhibitors should abolish α1(IV)NC1-induced caspase-8 and -9 activities in MRECs. Incubation of MRECs with z-IEPD-fmk and z-LEHD-fmk (specific caspase-8 and -9 inhibitors) showed the suppression of α1(IV)NC1-induced cellular apoptosis and the inhibition of caspase-8 and -9 activity (Figs. 5C 5D ; α1(IV)NC1+IEPD and α1(IV)NC1+LEDH). 
FACS Analysis of MREC Apoptosis by α1(IV)NC1
We further investigated the activation of apoptosis in MRECs incubated with and without α1(IV)NC1 with the use of FACS analysis. We quantified cellular apoptosis by measuring propidium iodide staining versus annexin V-FITC content in MRECs. MRECs incubated with control medium containing VEGF showed greater than 90% cell viability, whereas apoptosis was observed in less than 10% of total cell population (Fig. 6A) . The addition of α1(IV)NC1 under similar conditions decreased MREC viability to 26%, indicating that approximately 74% of cells underwent apoptosis (Fig. 6B) . Comprehensive statistical results of the observed apoptotic and nonapoptotic MRECs (incubated with and without α1(IV)NC1) are summarized in Figure 6C . Among the known caspases, caspase-3 is an important effector molecule for most cellular apoptosis. 26 To study whether caspase-3 could be activated by α1(IV)NC1, we incubated MRECs with α1(IV)NC1 for different time periods and observed the activation of caspase-3 in a time-dependent manner (Fig. 6D , lower blot). The time-dependent activation of caspase-3 band intensities is shown in Figure 6D(upper graph). 
Specific Apoptotic Effects of α1(IV)NC1 on p38-MAPK Signaling in Regulation of Bcl-2/Bcl-xL and Caspase-3/PARP Activation in MRECs
In this study we investigated whether α1(IV)NC1 played a role in the activation of caspase-3–mediated MREC apoptosis through the inactivation of p38-MAPK pathway. The p38-MAPK pathway is an important downstream target of FAK that is critical for the regulation of endothelial cell migration. 27 28 Mice deficient in FAK display severely impaired vasculogenesis and cell migration, whereas overexpression significantly increases the migratory capacity of cells through activation of the MAPK pathway. 29 30 These findings suggest that type IV collagen integrins/FAK/p38-MAPK pathway may play a major role in the regulation of matrix-mediated migration. Attachment of VEGF-treated MRECs to type IV collagen activated the FAK/p38 pathway (data not shown). Pretreatment of MRECs with α1(IV)NC1 did not inhibit the expression of FAK or p38-MAPK but significantly blocked the sustained phosphorylation of FAK and p38-MAPK (Figs. 7A 7C , top panels). In endothelial cells, VEGF-induced migration and cytoskeletal reorganization are mediated by ERKs-MAPK, whereas the expression of integrins and proteases is regulated by p38-MAPK. 31 32 In contrast, the incubation of MRPE cells with α1(IV)NC1 had no effect on the phosphorylation of FAK or p38-MAPK on type IV collagen matrix, suggesting that α1(IV)NC1-induced signaling is specific to endothelial cells (Figs. 7B 7D , upper panel). 
To understand whether the p38-MAPK pathway is involved in the proapoptotic activity of α1(IV)NC1, we examined changes in the expression of proapoptotic (Bax) and antiapoptotic (Bcl-2, Bcl-xL) proteins in MRECs treated with α1(IV)NC1. α1(IV)NC1-treated MRECs did not show any effect on Bax expression (data not shown), whereas Bcl-xL expression and Bcl-2 expression were significantly inhibited at 24 hours (Fig. 7E , lane 2 vs. lane 1). Earlier we reported that α1(IV)NC1 regulates HIF-1α expression dependent on p38-MAPK. 13 Here, we evaluated whether α1(IV)NC1-regulated Bcl-2 and Bcl-xL expressions were mediated through p38-MAPK. A specific p38-MAPK inhibitor, SB203580, should inhibit Bcl-2 and Bcl-xL expression; treatment of MRECs with SB203580 showed the inhibition of Bcl-2 and Bcl-xL, confirming that expression of these antiapoptotic molecules in MRECs by α1(IV)NC1 is dependent on p38-MAPK (Fig. 7E , lane 3). This is consistent with p38-MAPK–mediated endothelial cell apoptosis through the downregulation of Bcl-xL and Bcl-2. 33 Here, we also identified that activation of caspase-3 and PARP cleavage downstream of Bcl-2/Bcl-xL in MRECs treated with α1(IV)NC1 was p38-MAPK dependent (Fig. 7F , lane 2 vs. lane 1). In addition treatment of MRECs with p38-MAPK inhibitor (SB203580) showed the activation of caspase-3 and PARP cleavage, further confirming that inactivation of the p38-MAPK pathway by α1(IV)NC1 had an impact on MREC apoptosis (Fig. 7E , lane 3). 
Regulation of VEGF-Induced Neovascularization by α1(IV)NC1
We further evaluated the effects of baculovirus-expressed recombinant human α1(IV)NC1 on VEGF-mediated angiogenesis in vivo using the BMM plug assay in FVB/NJ mice. BMM plugs containing VEGF were used to assess the role of different doses of α1(IV)NC1 (0.5–1.0 μM) in the inhibition of VEGF-induced neovascularization. A 1.0-μM concentration of α1(IVNC1 significantly (nearly 88%) inhibited VEGF-induced neovascularization in the BMM plugs (Fig. 8A) . The number of blood vessels in the BMM plugs were as follows: VEGF + α1(IV)NC1 (0.5 μM), 17.45 ± 0.28; VEGF + α1(IV)NC1 (0.5 μM), 11.05 ± 0.15; controls, VEGF with and without collagen type IV, 30.0 ± 2.5 (Fig. 8B) . In contrast, the hemoglobin contents in the BMM plugs were as follows: VEGF + α1(IV)NC1 (0.5 μM) treated, 4.15 ± 0.25 g/dL (n = 6); VEGF + α1(IV)NC1 (1.0 μM) treated, 2.75 ± 0.16 g/dL (n = 6); hemoglobin content in VEGF with and without type IV collagen controls, 10.0 ± 1.2 g/dL (n = 6; Fig. 8C ). These results suggest that the administration of recombinant α1(IV)NC1 inhibits VEGF-mediated neovascularization in vivo. A schematic illustrates the distinct apoptotic signaling induced by α1(IV)NC1 in MREC apoptosis, presumably through the inhibition of FAK/p38-MAPK/Bcl-2/Bcl-xL, caspase-3 activation, and PARP cleavage (Fig. 8D)
Discussion
The NC1 domain generated from type IV collagen α1 chain α1(IV)NC1 was identified as an antiangiogenic molecule, and its activity was mediated through α1β1 integrin, specifically in proliferating endothelial cells. 13 34 However, it is critical to examine α1(IV)NC1 effects in several well-defined relevant in vitro model culture systems before assuming that this NC1 domain is universally antiangiogenic. In this study, we examined the effects of baculovirus-expressed human α1(IV)NC1 protein in in vitro and in vivo models of neovascularization. We demonstrated, for the first time, that α1(IV)NC1 inhibits VEGF-induced MREC proliferation, migration, and tube formation through the FAK/p38-MAPK pathway. In addition, we discovered the proapoptotic effect of α1(IV)NC1 mediation through the regulation of antiapoptotic Bcl-xL/Bcl-2 expression and activation of caspase-3/PARP cleavage through the inhibition of p38-MAPK signaling. This finding is consistent with results of previous studies demonstrating that antiangiogenic activity of α1(IV)NC1 is mediated through α1β1 integrin and inhibition of hypoxia-mediated signaling and apoptosis. 13 35 α1(IV)NC1 inhibited the phosphorylation of FAK and p38-MAPK when MRECs were plated on type IV collagen matrix. Similarly, we previously reported that another collagen NC1 domain, α3(IV)NC1, inhibits the phosphorylation of FAK on fibronectin matrix. 18 Understanding the mechanism(s) of action of these molecules is crucial for their therapeutic development and use. 
To further support the inhibition of angiogenesis observed in MRECs incubated with α1(IV)NC1 in culture, we assessed its activity in vivo. The intravenous administration of α1(IV)NC1 caused selective inhibition of endothelial cells growth in the VEGF-stimulated BMM, resulting in the suppression of neovascularization. Thus, α1(IV)NC1, which has previously been shown to suppress tumor angiogenesis in different mouse models, is also a strong antiangiogenic agent in MRECs and may be an effective therapeutic candidate for the treatment of neovascular diseases in the eye. 
Current evidence indicates that VEGF is a major angiogenic factor and plays a prominent role in ocular neovascularization. 36 VEGF is massively upregulated by hypoxia, and its levels are increased in the retina and vitreous of patients and in animal models of ischemic retinopathy. 37 38 39 VEGF is responsible for the growth of new blood vessels through the stimulation of endothelial cell proliferation and migration. 36 Increased expression of VEGF in transgenic mice stimulates neovascularization within the retina, and VEGF receptor antagonists inhibit retinal and choroidal neovascularization in many animal models. 40 41 Compared with VEGF expression, less evidence implicates basic fibroblast growth factor (FGF-2) in the development of ocular neovascularization. 42  
Recently, many researchers have reported that several antiangiogenic drugs (e.g., pegaptanib sodium [Macugen; Pfizer, New York, NY], RhuFab V2 [Lucentis; Genentech, South San Francisco, CA], tryptophanyl-tRNA synthetase (TrpRS), VEGF-TRAP, AdPEDF, AG-013958, bevacizumab [Avastin; Genentech], JSM6427) inhibit ocular neovascularization and prevent leakiness of retinal blood vessels in vivo by preventing the binding of VEGF to its receptors on endothelial cells. 6 43 44 45 46 47 48 A known endogenous angiogenesis inhibitor, α2(IV)NC1 (canstatin), was recently reported to inhibit laser-induced CNV by inducing apoptosis of endothelial cells. 49 Our findings suggest that α1(IV)NC1 may also be effective in the inhibition of choroidal neovascularization. This is further supported by earlier findings that type IV collagen–derived NC1 domains regulate angiogenesis in a number of in vitro and in vivo models. 11 Thus, this work not only supports our efforts in the development of α1(IV)NC1 as a potential candidate for the treatment of ocular diseases with a neovascular component, it also supports our ongoing oncology development efforts. 
 
Figure 1.
 
(A) MREC proliferation assay. Summary of relative [3H]-thymidine incorporation inhibition in MRECs on treatment with different concentrations of α1(IV)NC1 and α3(IV)NC1 compared with and without VEGF as controls. (B) MREC proliferation assays were also performed with methylene blue staining, treating cells with different concentrations of α1(IV)NC1 compared with and without VEGF as controls. All groups represent triplicate samples. (C, D) MREC and MRPE cell viability. MTT assay performed to evaluate MREC and MRPE cell viability after treatment with various concentrations of α1(IV)NC1. MRECs and MRPE cells grown with and without VEGF, shown as negative and positive controls. (AC) Results were significant; mean ± SEM of three independent experiments. *P < 0.001 and **P < 0.005 compared with VEGF treatment.
Figure 1.
 
(A) MREC proliferation assay. Summary of relative [3H]-thymidine incorporation inhibition in MRECs on treatment with different concentrations of α1(IV)NC1 and α3(IV)NC1 compared with and without VEGF as controls. (B) MREC proliferation assays were also performed with methylene blue staining, treating cells with different concentrations of α1(IV)NC1 compared with and without VEGF as controls. All groups represent triplicate samples. (C, D) MREC and MRPE cell viability. MTT assay performed to evaluate MREC and MRPE cell viability after treatment with various concentrations of α1(IV)NC1. MRECs and MRPE cells grown with and without VEGF, shown as negative and positive controls. (AC) Results were significant; mean ± SEM of three independent experiments. *P < 0.001 and **P < 0.005 compared with VEGF treatment.
Figure 2.
 
(A) MREC migration assay. Numbers of MRECs that migrated in VEGF with and without α1(IV)NC1 were evaluated with light microscopy, and representative fields are shown at 100× magnification. MRECs on the underside of a Boyden chamber membrane are shown. (B) MREC migration assessment. Average number of MRECs migrated in four different wells in each condition (n = 4); mean ± SEM of three independent experiments. *P < 0.001 compared with VEGF treatment.
Figure 2.
 
(A) MREC migration assay. Numbers of MRECs that migrated in VEGF with and without α1(IV)NC1 were evaluated with light microscopy, and representative fields are shown at 100× magnification. MRECs on the underside of a Boyden chamber membrane are shown. (B) MREC migration assessment. Average number of MRECs migrated in four different wells in each condition (n = 4); mean ± SEM of three independent experiments. *P < 0.001 compared with VEGF treatment.
Figure 3.
 
(A) Tube formation assay. MRECs were plated on BMM-coated plates in retinal endothelial cell medium as control or with 0.25 to 1.0 μM α1(IV)NC1 protein. Tube formation was evaluated with light microscopy, and representative fields are shown at 100× magnification. (B) Tube formation assessments. Average number of tubes in two wells in each condition and mean ± SEM of three independent experiments (n = 6). *P < 0.001 compared with control.
Figure 3.
 
(A) Tube formation assay. MRECs were plated on BMM-coated plates in retinal endothelial cell medium as control or with 0.25 to 1.0 μM α1(IV)NC1 protein. Tube formation was evaluated with light microscopy, and representative fields are shown at 100× magnification. (B) Tube formation assessments. Average number of tubes in two wells in each condition and mean ± SEM of three independent experiments (n = 6). *P < 0.001 compared with control.
Figure 4.
 
Impaired migration and proliferation of MRECs incubated with α1(IV)NC1 in a scratch wound assay. (A) MRECs were cultured to 80% confluence in 24-well plates in serum-containing medium. Wounds were created in the MREC monolayer using a sterile pipette tip. Photographs were taken immediately and later at indicated time intervals after the wounds were created. Data are from one representative experiment. Similar results were obtained in two independent experiments. (B) Bar diagram represents quantification of the wound-healing assessment from duplicate wells of three independent experiments (n = 6). *P < 0.001 for control versus α1(IV)NC1-treated MRECs.
Figure 4.
 
Impaired migration and proliferation of MRECs incubated with α1(IV)NC1 in a scratch wound assay. (A) MRECs were cultured to 80% confluence in 24-well plates in serum-containing medium. Wounds were created in the MREC monolayer using a sterile pipette tip. Photographs were taken immediately and later at indicated time intervals after the wounds were created. Data are from one representative experiment. Similar results were obtained in two independent experiments. (B) Bar diagram represents quantification of the wound-healing assessment from duplicate wells of three independent experiments (n = 6). *P < 0.001 for control versus α1(IV)NC1-treated MRECs.
Figure 5.
 
Caspase-3 activation. (A, B) MRECs and MRPE cells incubated with and without α1(IV)NC1 and cytosolic extracts were analyzed for caspase-3 activity. DEVD-fmk and TNF-α were used as positive control. (A, B) Results were significant; mean ± SEM of three independent experiments is shown. *P < 0.001 compared with control; **P < 0.001 compared with α1(IV)NC1 treatment. (C, D) Activation of caspase-8 and -9 by α1(IV)NC1 at 24 hours as measured by cleavage of specific substrates IEDP-AMC and LEDH-AFC. Staurosporine was used as a positive control. Results are shown as mean ± SEM from three independent experiments. *P < 0.005 compared with control; **P < 0.005 compared with α1(IV)NC1 treatment.
Figure 5.
 
Caspase-3 activation. (A, B) MRECs and MRPE cells incubated with and without α1(IV)NC1 and cytosolic extracts were analyzed for caspase-3 activity. DEVD-fmk and TNF-α were used as positive control. (A, B) Results were significant; mean ± SEM of three independent experiments is shown. *P < 0.001 compared with control; **P < 0.001 compared with α1(IV)NC1 treatment. (C, D) Activation of caspase-8 and -9 by α1(IV)NC1 at 24 hours as measured by cleavage of specific substrates IEDP-AMC and LEDH-AFC. Staurosporine was used as a positive control. Results are shown as mean ± SEM from three independent experiments. *P < 0.005 compared with control; **P < 0.005 compared with α1(IV)NC1 treatment.
Figure 6.
 
MRECs were treated with and without α1(IV)NC1, stained with annexin V-FITC and PI, and analyzed by flow cytometry. (A, B) Dot blots shown with and without α1(IV)NC1-treated MREC survival that was measured by annexin V and PI cell populations (arrows). (C) Bar graph indicates percentages of annexin V and PI cells after they were grown on type IV collagen-coated tissue cultures plates for 24 hours. Similar results were also obtained in two other independent experiments conducted in triplicate. *P < 0.005 α1(IV)NC1-treated apoptotic versus nonapoptotic MRECs. (D, top) Control and α1(IV)NC1-treated MRECs were lysed, and caspase-3 activity was determined with caspase-3 specific antibody (upper and lower arrows in the immunoblot indicate pro-caspase-3 and active caspase-3). Bottom: relative densities of pixels and area of activated caspase-3 bands were determined with ImageJ software (developed by Wayne Rasband, National Institutes of Health, Bethesda, MD; available at http://rsb.info.nih.gov/ij/index.html).
Figure 6.
 
MRECs were treated with and without α1(IV)NC1, stained with annexin V-FITC and PI, and analyzed by flow cytometry. (A, B) Dot blots shown with and without α1(IV)NC1-treated MREC survival that was measured by annexin V and PI cell populations (arrows). (C) Bar graph indicates percentages of annexin V and PI cells after they were grown on type IV collagen-coated tissue cultures plates for 24 hours. Similar results were also obtained in two other independent experiments conducted in triplicate. *P < 0.005 α1(IV)NC1-treated apoptotic versus nonapoptotic MRECs. (D, top) Control and α1(IV)NC1-treated MRECs were lysed, and caspase-3 activity was determined with caspase-3 specific antibody (upper and lower arrows in the immunoblot indicate pro-caspase-3 and active caspase-3). Bottom: relative densities of pixels and area of activated caspase-3 bands were determined with ImageJ software (developed by Wayne Rasband, National Institutes of Health, Bethesda, MD; available at http://rsb.info.nih.gov/ij/index.html).
Figure 7.
 
(A, B) FAK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-FAK indicate that VEGF-mediated phosphorylation of FAK (p-FAK) was inhibited in MRECs incubated with α1(IV)NC1 but not in MRPE cells (A, B, top) or total FAK proteins (A, B, bottom). (C, D) p38-MAPK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-p38 indicate that VEGF-mediated sustained phosphorylation of p38 (p-p38) was inhibited by incubation with α1(IV)NC1 in MRECs but not in MRPE cells (C, D, top) and total p38-MAPK (C, D, bottom). (E, F) Effect of α1(IV)NC1 on p38 MAPK-dependent regulation of Bcl-2/Bcl-xL and activation of caspase-3/PARP. VEGF-stimulated MRECs treated with PBS (lane 1, control), 1 μM α1(IV) NC1 (lane 2), 20 μM SB203580, a specific p38-MAPK inhibitor (lane 3), and 80 ng/mL TNF-α (lane 4) for 24 hours. Total cells were collected and lysed for 30 minutes in ice-cold RIPA lysis buffer, and 25 μg total cytosolic extract per lane was separated and immunoblotted with primary antibodies against signaling molecules in the mitochondrial apoptotic pathway.
Figure 7.
 
(A, B) FAK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-FAK indicate that VEGF-mediated phosphorylation of FAK (p-FAK) was inhibited in MRECs incubated with α1(IV)NC1 but not in MRPE cells (A, B, top) or total FAK proteins (A, B, bottom). (C, D) p38-MAPK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-p38 indicate that VEGF-mediated sustained phosphorylation of p38 (p-p38) was inhibited by incubation with α1(IV)NC1 in MRECs but not in MRPE cells (C, D, top) and total p38-MAPK (C, D, bottom). (E, F) Effect of α1(IV)NC1 on p38 MAPK-dependent regulation of Bcl-2/Bcl-xL and activation of caspase-3/PARP. VEGF-stimulated MRECs treated with PBS (lane 1, control), 1 μM α1(IV) NC1 (lane 2), 20 μM SB203580, a specific p38-MAPK inhibitor (lane 3), and 80 ng/mL TNF-α (lane 4) for 24 hours. Total cells were collected and lysed for 30 minutes in ice-cold RIPA lysis buffer, and 25 μg total cytosolic extract per lane was separated and immunoblotted with primary antibodies against signaling molecules in the mitochondrial apoptotic pathway.
Figure 8.
 
Regulation of VEGF-induced neovascularization of BMM implants in mice. (A) FVB/NJ mice. Left to right: different conditions of BMM are shown. Arrows: blood vessels. E, endothelial cells; M, BMM; SM, smooth muscle cells. Scale bar, 50 μm. (B, C) Number of blood vessels and hemoglobin (Hb) content quantification from (A). Mean ± SEM. *P < 0.01 compared with VEGF and without α1(IV)NC1 (0.5 μM); **P < 0.005 compared with VEGF and without α1(IV)NC1 (1 μM). Blood vessels in the BMM plug were counted in 10 fields at 200× magnification (n = 6). (D) Schematic illustrates distinct apoptotic signaling induced by α1(IV)NC1 in MRECs. α1(IV)NC1 binds to α1β1 integrin and initiates two apoptotic pathways, involving the activation of caspase-9 and -8 and leading to the activation of caspase-3 and PARP cleavage. (1) α1(IV)NC1 activates caspase-3 directly through the inhibition of FAK/p38-MAPK/Bcl-2/Bcl-xL and the activation of caspase-9. (2) Integrin cross-talk with Fas through the mitochondrial pathway led to the activation of caspase-8 and -3.
Figure 8.
 
Regulation of VEGF-induced neovascularization of BMM implants in mice. (A) FVB/NJ mice. Left to right: different conditions of BMM are shown. Arrows: blood vessels. E, endothelial cells; M, BMM; SM, smooth muscle cells. Scale bar, 50 μm. (B, C) Number of blood vessels and hemoglobin (Hb) content quantification from (A). Mean ± SEM. *P < 0.01 compared with VEGF and without α1(IV)NC1 (0.5 μM); **P < 0.005 compared with VEGF and without α1(IV)NC1 (1 μM). Blood vessels in the BMM plug were counted in 10 fields at 200× magnification (n = 6). (D) Schematic illustrates distinct apoptotic signaling induced by α1(IV)NC1 in MRECs. α1(IV)NC1 binds to α1β1 integrin and initiates two apoptotic pathways, involving the activation of caspase-9 and -8 and leading to the activation of caspase-3 and PARP cleavage. (1) α1(IV)NC1 activates caspase-3 directly through the inhibition of FAK/p38-MAPK/Bcl-2/Bcl-xL and the activation of caspase-9. (2) Integrin cross-talk with Fas through the mitochondrial pathway led to the activation of caspase-8 and -3.
FolkmanJ. Angiogenesis. Annu Rev Med. 2006;57:1–18. [CrossRef] [PubMed]
GarnerA. Ocular angiogenesis. Int Rev Exp Pathol. 1986;28:249–306. [PubMed]
JampolLM, EbroonDA, GoldbaumMH. Peripheral proliferative retinopathies: an update on angiogenesis, etiologies and management. Surv Ophthalmol. 1994;38:519–540. [CrossRef] [PubMed]
FolkmanJ. Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat Med. 1995;1:27–31. [CrossRef] [PubMed]
TakahashiH, TamakiY, IshiiN, et al. Identification of a novel vascular endothelial growth factor receptor 2 inhibitor and its effect for choroidal neovascularization in vivo. Curr Eye Res. 2008;33:1002–1010. [CrossRef] [PubMed]
DeisslerHL, LangGE. [Effect of VEGF165 and the VEGF aptamer pegaptanib (Macugen) on the protein composition of tight junctions in microvascular endothelial cells of the retina]. Klin Monatsbl Augenheilkd. 2008;225:863–867. [CrossRef] [PubMed]
PaulssonM. Basement membrane proteins: structure, assembly, and cellular interactions. Crit Rev Biochem Mol Biol. 1992;27:93–127. [CrossRef] [PubMed]
IngberD, FolkmanJ. Inhibition of angiogenesis through modulation of collagen metabolism. Lab Invest. 1988;59:44–51. [PubMed]
CarmelietP, JainRK. Angiogenesis in cancer and other diseases. Nature. 2000;407:249–257. [CrossRef] [PubMed]
MaragoudakisME, MissirlisE, KarakiulakisGD, SarmonicaM, BastakisM, TsopanoglouN. Basement membrane biosynthesis as a target for developing inhibitors of angiogenesis with anti-tumor properties. Kidney Int. 1993;43:147–150. [CrossRef] [PubMed]
PedchenkoV, ZentR, HudsonBG. Alpha (v) beta3 and alpha (v) beta5 integrins bind both the proximal RGD site and non-RGD motifs within noncollagenous (NC1) domain of the alpha3 chain of type IV collagen: implication for the mechanism of endothelia cell adhesion. J Biol Chem. 2004;279:2772–2780. [CrossRef] [PubMed]
ColoradoPC, TorreA, KamphausG, et al. Anti-angiogenic cues from vascular basement membrane collagen. Cancer Res. 2000;60:2520–2526. [PubMed]
SudhakarA, NybergP, KeshamouniVG, et al. Human alpha1 type IV collagen NC1 domain exhibits distinct antiangiogenic activity mediated by alpha1beta1 integrin. J Clin Invest. 2005;115:2801–2810. [CrossRef] [PubMed]
SuX, SorensonCM, SheibaniN. Isolation and characterization of murine retinal endothelial cells. Mol Vis. 2003;9:171–178. [PubMed]
BorzaCM, PozziA, BorzaDB, et al. Integrin alpha3beta1, a novel receptor for alpha3(IV) noncollagenous domain and a trans-dominant inhibitor for integrin alphavbeta3. J Biol Chem. 2006;281:20932–20939. [CrossRef] [PubMed]
BoosaniCS, SudhakarA. Cloning, purification, and characterization of a non-collagenous anti-angiogenic protein domain from human alpha1 type IV collagen expressed in Sf9 cells. Protein Expr Purif. 2006;49:211–218. [CrossRef] [PubMed]
ChowJ, OgunsholaO, FanSY, LiY, MentLR, MadriJA. Astrocyte-derived VEGF mediates survival and tube stabilization of hypoxic brain microvascular endothelial cells in vitro. Brain Res Dev Brain Res. 2001;130:123–132. [CrossRef] [PubMed]
BoosaniCS, MannamAP, CosgroveD, et al. Regulation of COX-2 mediated signaling by α3 type IV noncollagenous domain in tumor angiogenesis. Blood. 2007;110:1168–1177. [CrossRef] [PubMed]
MunugalavadlaV, BorneoJ, IngramDA, KapurR. p85alpha subunit of class IA PI-3 kinase is crucial for macrophage growth and migration. Blood. 2005;106:103–109. [CrossRef] [PubMed]
KohlerC, OrreniusS, ZhivotovskyB. Evaluation of caspase activity in apoptotic cells. J Immunol Methods. 2002;265:97–110. [CrossRef] [PubMed]
AoudjitF, VuoriK. Integrin signaling inhibits paclitaxel-induced apoptosis in breast cancer cells. Oncogene. 2001;20:4995–5004. [CrossRef] [PubMed]
BagleyRG, Walter-YohrlingJ, CaoX, et al. Endothelial precursor cells as a model of tumor endothelium: characterization and comparison with mature endothelial cells. Cancer Res. 2003;63:5866–5873. [PubMed]
McCawleyLJ, MatrisianLM. Tumor progression: defining the soil round the tumor seed. Curr Biol. 2001;11:R25–R27. [CrossRef] [PubMed]
HuangQ, SheibaniN. High glucose promotes retinal endothelial cell migration through activation of Src, PI3K/Akt1/eNOS, and ERKs. Am J Physiol Cell Physiol. 2008;295:C1647–C1657. [CrossRef] [PubMed]
FolkmanJ, HaudenschildC. Angiogenesis by capillary endothelial cells in culture. Trans Ophthalmol Soc UK. 1980;100:346–353. [PubMed]
NunezG, BenedictMA, HuY, InoharaN. Caspases: the proteases of the apoptotic pathway. Oncogene. 1998;17:3237–3245. [PubMed]
ClarkEA, BruggeJS. Integrins and signal transduction pathways: the road taken. Science. 1995;268:233–239. [CrossRef] [PubMed]
RobertsMS, WoodsAJ, ShawPE, NormanJC. ERK1 associates with alpha vbeta 3 integrin and regulates cell spreading on vitronectin. J Biol Chem. 2003;278:1975–1985. [CrossRef] [PubMed]
FurutaY, IlicD, KanazawaS, TakedaN, YamamotoT, AizawaS. Mesodermal defect in late phase of gastrulation by a targeted mutation of focal adhesion kinase, FAK. Oncogene. 1995;11:1989–1995. [PubMed]
CaryLA, HanDC, PolteTR, HanksSK, GuanJL. Identification of p130Cas as a mediator of focal adhesion kinase- promoted cell migration. J Cell Biol. 1998;140:211–221. [CrossRef] [PubMed]
BeckerPM, VerinAD, BoothMA, LiuF, BirukovaA, GarciaJG. Differential regulation of diverse physiological responses to VEGF in pulmonary endothelial cells. Am J Physiol Lung Cell Mol Physiol. 2001;281:L1500–L1511. [PubMed]
KuoCJ, LaMontagneKR, Jr, Garcia-CardenaG, et al. Oligomerization-dependent regulation of motility and morphogenesis by the collagen XVIII NC1/endostatin domain. J Cell Biol. 2001;152:1233–1246. [CrossRef] [PubMed]
GretheS, AresMP, AnderssonT, Porn-AresMI. p38 MAPK mediates TNF-induced apoptosis in endothelial cells via phosphorylation and downregulation of Bcl-x(L). Exp Cell Res. 2004;298:632–642. [CrossRef] [PubMed]
SudhakarA, BoosaniCS. Signaling mechanisms of endogenous angiogenesis inhibitors derived from type IV collagen. Gene Regulation Syst Biol. 2007;1:217–226.
NybergP, XieL, SugimotoH, et al. Characterization of the anti-angiogenic properties of arrestin, an α1β1 integrin-dependent collagen-derived tumor suppressor. Exp Cell Res. 2008;314:3292–3305. [CrossRef] [PubMed]
SmithLE, KopchickJJ, ChenW, et al. Essential role of growth hormone in ischemia-induced retinal neovascularization. Science. 1997;276:1706–1709. [CrossRef] [PubMed]
ShweikiD, ItinA, SofferD, KeshetE. Vascular endothelial growth factor induced by hypoxia may mediate hypoxia-initiated angiogenesis. Nature. 1992;359:843–845. [CrossRef] [PubMed]
AdamisAP, MillerJW, BernalMT, et al. Increased vascular endothelial growth factor levels in the vitreous of eyes with proliferative diabetic retinopathy. Am J Ophthalmol. 1994;118:445–450. [CrossRef] [PubMed]
PierceEA, AveryRL, FoleyED, AielloLP, SmithLE. Vascular endothelial growth factor/vascular permeability factor expression in a mouse model of retinal neovascularization. Proc Natl Acad Sci U S A. 1995;92:905–909. [CrossRef] [PubMed]
OkamotoN, TobeT, HackettSF, et al. Transgenic mice with increased expression of vascular endothelial growth factor in the retina: a new model of intraretinal and subretinal neovascularization. Am J Pathol. 1997;151:281–291. [PubMed]
TobeT, OkamotoN, VinoresMA, et al. Evolution of neovascularization in mice with overexpression of vascular endothelial growth factor in photoreceptors. Invest Ophthalmol Vis Sci. 1998;39:180–188. [PubMed]
TobeT, OrtegaS, LunaJD, et al. Targeted disruption of the FGF2 gene does not prevent choroidal neovascularization in a murine model. Am J Pathol. 1998;153:1641–1646. [CrossRef] [PubMed]
MantelI, ZografosL, AmbresinA. Early clinical experience with ranibizumab for occult and minimally classic neovascular membranes in age-related macular degeneration. Ophthalmologica. 2008;222:321–323. [CrossRef] [PubMed]
KonstantinidisL, MantelI, PournarasJA, ZografosL, AmbresinA. Intravitreal ranibizumab (Lucentis) for the treatment of myopic choroidal neovascularization. Graefes Arch Clin Exp Ophthalmol. 2009;247:311–318. [CrossRef] [PubMed]
SiemannDW, ShiW. Dual targeting of tumor vasculature: combining Avastin and vascular disrupting agents (CA4P or OXi4503). Anticancer Res. 2008;28:2027–2031. [PubMed]
NguyenQD, ShahSM, HafizG, et al. A phase I trial of an IV-administered vascular endothelial growth factor trap for treatment in patients with choroidal neovascularization due to age-related macular degeneration. Ophthalmology. 2006;113:1522.e1521–1522.e1514.
CampochiaroPA, NguyenQD, ShahSM, et al. Adenoviral vector-delivered pigment epithelium-derived factor for neovascular age-related macular degeneration: results of a phase I clinical trial. Hum Gene Ther. 2006;17:167–176. [CrossRef] [PubMed]
LuF, AdelmanRA. Are intravitreal bevacizumab and ranibizumab effective in a rat model of choroidal neovascularization?. Graefes Arch Clin Exp Ophthalmol. 2009;247:171–177. [CrossRef] [PubMed]
LimaESR, KachiS, AkiyamaH, et al. Recombinant non-collagenous domain of alpha2(IV) collagen causes involution of choroidal neovascularization by inducing apoptosis. J Cell Physiol. 2006;208:161–166. [CrossRef] [PubMed]
Figure 1.
 
(A) MREC proliferation assay. Summary of relative [3H]-thymidine incorporation inhibition in MRECs on treatment with different concentrations of α1(IV)NC1 and α3(IV)NC1 compared with and without VEGF as controls. (B) MREC proliferation assays were also performed with methylene blue staining, treating cells with different concentrations of α1(IV)NC1 compared with and without VEGF as controls. All groups represent triplicate samples. (C, D) MREC and MRPE cell viability. MTT assay performed to evaluate MREC and MRPE cell viability after treatment with various concentrations of α1(IV)NC1. MRECs and MRPE cells grown with and without VEGF, shown as negative and positive controls. (AC) Results were significant; mean ± SEM of three independent experiments. *P < 0.001 and **P < 0.005 compared with VEGF treatment.
Figure 1.
 
(A) MREC proliferation assay. Summary of relative [3H]-thymidine incorporation inhibition in MRECs on treatment with different concentrations of α1(IV)NC1 and α3(IV)NC1 compared with and without VEGF as controls. (B) MREC proliferation assays were also performed with methylene blue staining, treating cells with different concentrations of α1(IV)NC1 compared with and without VEGF as controls. All groups represent triplicate samples. (C, D) MREC and MRPE cell viability. MTT assay performed to evaluate MREC and MRPE cell viability after treatment with various concentrations of α1(IV)NC1. MRECs and MRPE cells grown with and without VEGF, shown as negative and positive controls. (AC) Results were significant; mean ± SEM of three independent experiments. *P < 0.001 and **P < 0.005 compared with VEGF treatment.
Figure 2.
 
(A) MREC migration assay. Numbers of MRECs that migrated in VEGF with and without α1(IV)NC1 were evaluated with light microscopy, and representative fields are shown at 100× magnification. MRECs on the underside of a Boyden chamber membrane are shown. (B) MREC migration assessment. Average number of MRECs migrated in four different wells in each condition (n = 4); mean ± SEM of three independent experiments. *P < 0.001 compared with VEGF treatment.
Figure 2.
 
(A) MREC migration assay. Numbers of MRECs that migrated in VEGF with and without α1(IV)NC1 were evaluated with light microscopy, and representative fields are shown at 100× magnification. MRECs on the underside of a Boyden chamber membrane are shown. (B) MREC migration assessment. Average number of MRECs migrated in four different wells in each condition (n = 4); mean ± SEM of three independent experiments. *P < 0.001 compared with VEGF treatment.
Figure 3.
 
(A) Tube formation assay. MRECs were plated on BMM-coated plates in retinal endothelial cell medium as control or with 0.25 to 1.0 μM α1(IV)NC1 protein. Tube formation was evaluated with light microscopy, and representative fields are shown at 100× magnification. (B) Tube formation assessments. Average number of tubes in two wells in each condition and mean ± SEM of three independent experiments (n = 6). *P < 0.001 compared with control.
Figure 3.
 
(A) Tube formation assay. MRECs were plated on BMM-coated plates in retinal endothelial cell medium as control or with 0.25 to 1.0 μM α1(IV)NC1 protein. Tube formation was evaluated with light microscopy, and representative fields are shown at 100× magnification. (B) Tube formation assessments. Average number of tubes in two wells in each condition and mean ± SEM of three independent experiments (n = 6). *P < 0.001 compared with control.
Figure 4.
 
Impaired migration and proliferation of MRECs incubated with α1(IV)NC1 in a scratch wound assay. (A) MRECs were cultured to 80% confluence in 24-well plates in serum-containing medium. Wounds were created in the MREC monolayer using a sterile pipette tip. Photographs were taken immediately and later at indicated time intervals after the wounds were created. Data are from one representative experiment. Similar results were obtained in two independent experiments. (B) Bar diagram represents quantification of the wound-healing assessment from duplicate wells of three independent experiments (n = 6). *P < 0.001 for control versus α1(IV)NC1-treated MRECs.
Figure 4.
 
Impaired migration and proliferation of MRECs incubated with α1(IV)NC1 in a scratch wound assay. (A) MRECs were cultured to 80% confluence in 24-well plates in serum-containing medium. Wounds were created in the MREC monolayer using a sterile pipette tip. Photographs were taken immediately and later at indicated time intervals after the wounds were created. Data are from one representative experiment. Similar results were obtained in two independent experiments. (B) Bar diagram represents quantification of the wound-healing assessment from duplicate wells of three independent experiments (n = 6). *P < 0.001 for control versus α1(IV)NC1-treated MRECs.
Figure 5.
 
Caspase-3 activation. (A, B) MRECs and MRPE cells incubated with and without α1(IV)NC1 and cytosolic extracts were analyzed for caspase-3 activity. DEVD-fmk and TNF-α were used as positive control. (A, B) Results were significant; mean ± SEM of three independent experiments is shown. *P < 0.001 compared with control; **P < 0.001 compared with α1(IV)NC1 treatment. (C, D) Activation of caspase-8 and -9 by α1(IV)NC1 at 24 hours as measured by cleavage of specific substrates IEDP-AMC and LEDH-AFC. Staurosporine was used as a positive control. Results are shown as mean ± SEM from three independent experiments. *P < 0.005 compared with control; **P < 0.005 compared with α1(IV)NC1 treatment.
Figure 5.
 
Caspase-3 activation. (A, B) MRECs and MRPE cells incubated with and without α1(IV)NC1 and cytosolic extracts were analyzed for caspase-3 activity. DEVD-fmk and TNF-α were used as positive control. (A, B) Results were significant; mean ± SEM of three independent experiments is shown. *P < 0.001 compared with control; **P < 0.001 compared with α1(IV)NC1 treatment. (C, D) Activation of caspase-8 and -9 by α1(IV)NC1 at 24 hours as measured by cleavage of specific substrates IEDP-AMC and LEDH-AFC. Staurosporine was used as a positive control. Results are shown as mean ± SEM from three independent experiments. *P < 0.005 compared with control; **P < 0.005 compared with α1(IV)NC1 treatment.
Figure 6.
 
MRECs were treated with and without α1(IV)NC1, stained with annexin V-FITC and PI, and analyzed by flow cytometry. (A, B) Dot blots shown with and without α1(IV)NC1-treated MREC survival that was measured by annexin V and PI cell populations (arrows). (C) Bar graph indicates percentages of annexin V and PI cells after they were grown on type IV collagen-coated tissue cultures plates for 24 hours. Similar results were also obtained in two other independent experiments conducted in triplicate. *P < 0.005 α1(IV)NC1-treated apoptotic versus nonapoptotic MRECs. (D, top) Control and α1(IV)NC1-treated MRECs were lysed, and caspase-3 activity was determined with caspase-3 specific antibody (upper and lower arrows in the immunoblot indicate pro-caspase-3 and active caspase-3). Bottom: relative densities of pixels and area of activated caspase-3 bands were determined with ImageJ software (developed by Wayne Rasband, National Institutes of Health, Bethesda, MD; available at http://rsb.info.nih.gov/ij/index.html).
Figure 6.
 
MRECs were treated with and without α1(IV)NC1, stained with annexin V-FITC and PI, and analyzed by flow cytometry. (A, B) Dot blots shown with and without α1(IV)NC1-treated MREC survival that was measured by annexin V and PI cell populations (arrows). (C) Bar graph indicates percentages of annexin V and PI cells after they were grown on type IV collagen-coated tissue cultures plates for 24 hours. Similar results were also obtained in two other independent experiments conducted in triplicate. *P < 0.005 α1(IV)NC1-treated apoptotic versus nonapoptotic MRECs. (D, top) Control and α1(IV)NC1-treated MRECs were lysed, and caspase-3 activity was determined with caspase-3 specific antibody (upper and lower arrows in the immunoblot indicate pro-caspase-3 and active caspase-3). Bottom: relative densities of pixels and area of activated caspase-3 bands were determined with ImageJ software (developed by Wayne Rasband, National Institutes of Health, Bethesda, MD; available at http://rsb.info.nih.gov/ij/index.html).
Figure 7.
 
(A, B) FAK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-FAK indicate that VEGF-mediated phosphorylation of FAK (p-FAK) was inhibited in MRECs incubated with α1(IV)NC1 but not in MRPE cells (A, B, top) or total FAK proteins (A, B, bottom). (C, D) p38-MAPK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-p38 indicate that VEGF-mediated sustained phosphorylation of p38 (p-p38) was inhibited by incubation with α1(IV)NC1 in MRECs but not in MRPE cells (C, D, top) and total p38-MAPK (C, D, bottom). (E, F) Effect of α1(IV)NC1 on p38 MAPK-dependent regulation of Bcl-2/Bcl-xL and activation of caspase-3/PARP. VEGF-stimulated MRECs treated with PBS (lane 1, control), 1 μM α1(IV) NC1 (lane 2), 20 μM SB203580, a specific p38-MAPK inhibitor (lane 3), and 80 ng/mL TNF-α (lane 4) for 24 hours. Total cells were collected and lysed for 30 minutes in ice-cold RIPA lysis buffer, and 25 μg total cytosolic extract per lane was separated and immunoblotted with primary antibodies against signaling molecules in the mitochondrial apoptotic pathway.
Figure 7.
 
(A, B) FAK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-FAK indicate that VEGF-mediated phosphorylation of FAK (p-FAK) was inhibited in MRECs incubated with α1(IV)NC1 but not in MRPE cells (A, B, top) or total FAK proteins (A, B, bottom). (C, D) p38-MAPK phosphorylation in MRECs and MRPE cells. Immunoblots for phospho-p38 indicate that VEGF-mediated sustained phosphorylation of p38 (p-p38) was inhibited by incubation with α1(IV)NC1 in MRECs but not in MRPE cells (C, D, top) and total p38-MAPK (C, D, bottom). (E, F) Effect of α1(IV)NC1 on p38 MAPK-dependent regulation of Bcl-2/Bcl-xL and activation of caspase-3/PARP. VEGF-stimulated MRECs treated with PBS (lane 1, control), 1 μM α1(IV) NC1 (lane 2), 20 μM SB203580, a specific p38-MAPK inhibitor (lane 3), and 80 ng/mL TNF-α (lane 4) for 24 hours. Total cells were collected and lysed for 30 minutes in ice-cold RIPA lysis buffer, and 25 μg total cytosolic extract per lane was separated and immunoblotted with primary antibodies against signaling molecules in the mitochondrial apoptotic pathway.
Figure 8.
 
Regulation of VEGF-induced neovascularization of BMM implants in mice. (A) FVB/NJ mice. Left to right: different conditions of BMM are shown. Arrows: blood vessels. E, endothelial cells; M, BMM; SM, smooth muscle cells. Scale bar, 50 μm. (B, C) Number of blood vessels and hemoglobin (Hb) content quantification from (A). Mean ± SEM. *P < 0.01 compared with VEGF and without α1(IV)NC1 (0.5 μM); **P < 0.005 compared with VEGF and without α1(IV)NC1 (1 μM). Blood vessels in the BMM plug were counted in 10 fields at 200× magnification (n = 6). (D) Schematic illustrates distinct apoptotic signaling induced by α1(IV)NC1 in MRECs. α1(IV)NC1 binds to α1β1 integrin and initiates two apoptotic pathways, involving the activation of caspase-9 and -8 and leading to the activation of caspase-3 and PARP cleavage. (1) α1(IV)NC1 activates caspase-3 directly through the inhibition of FAK/p38-MAPK/Bcl-2/Bcl-xL and the activation of caspase-9. (2) Integrin cross-talk with Fas through the mitochondrial pathway led to the activation of caspase-8 and -3.
Figure 8.
 
Regulation of VEGF-induced neovascularization of BMM implants in mice. (A) FVB/NJ mice. Left to right: different conditions of BMM are shown. Arrows: blood vessels. E, endothelial cells; M, BMM; SM, smooth muscle cells. Scale bar, 50 μm. (B, C) Number of blood vessels and hemoglobin (Hb) content quantification from (A). Mean ± SEM. *P < 0.01 compared with VEGF and without α1(IV)NC1 (0.5 μM); **P < 0.005 compared with VEGF and without α1(IV)NC1 (1 μM). Blood vessels in the BMM plug were counted in 10 fields at 200× magnification (n = 6). (D) Schematic illustrates distinct apoptotic signaling induced by α1(IV)NC1 in MRECs. α1(IV)NC1 binds to α1β1 integrin and initiates two apoptotic pathways, involving the activation of caspase-9 and -8 and leading to the activation of caspase-3 and PARP cleavage. (1) α1(IV)NC1 activates caspase-3 directly through the inhibition of FAK/p38-MAPK/Bcl-2/Bcl-xL and the activation of caspase-9. (2) Integrin cross-talk with Fas through the mitochondrial pathway led to the activation of caspase-8 and -3.
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