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[Cancer Research 64, 3679-3686, May 15, 2004]
© 2004 American Association for Cancer Research

Endocrinology

Endothelial Cell Surface ATP Synthase-Triggered Caspase-Apoptotic Pathway Is Essential for K1-5-Induced Antiangiogenesis

Niina Veitonmäki1, Renhai Cao1, Lin-Hua Wu3, Tammy L. Moser4, Bo Li4, Salvatore V. Pizzo4, Boris Zhivotovsky2 and Yihai Cao1

1 Microbiology and Tumor Biology Center and 2 Institute of Environmental Medicine, Karolinska Institutet, Stockholm, Sweden; 3 Department of Biochemistry, National Cheng Kung University Medical College, Taiwan, Republic of China; and 4 Department of Pathology and Duke University School of Nursing, Duke University Medical Center, Durham, North Carolina


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
We have recently reported the identification of kringle 1-5 (K1-5) of plasminogen as a potent and specific inhibitor of angiogenesis and tumor growth. Here, we show that K1-5 bound to endothelial cell surface ATP synthase and triggered caspase-mediated endothelial cell apoptosis. Induction of endothelial apoptosis involved sequential activation of caspases-8, -9, and -3. Administration of neutralizing antibodies directed against the {alpha}- and ß-subunits of ATP synthase to endothelial cells attenuated activation of these caspases. Furthermore, inhibitors of caspases-3, -8, and -9 also remarkably blocked K1-5-induced endothelial cell apoptosis and antiangiogenic responses. In a mouse tumor model, we show that caspase-3 inhibitors abolished the antitumor activity of K1-5 by protecting the tumor vasculature undergoing apoptosis. These results suggest that the specificity of the antiendothelial effect of K1-5 is attributable, at least in part, to its interaction with the endothelial cell surface ATP synthase and that the caspase-mediated endothelial apoptosis is essential for the angiostatic activity of K1-5. Thus, our findings provide a mechanistic insight with respect to the angiostatic action and signaling pathway of K1-5 and angiostatin.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Endogenous angiogenesis inhibitors produced by various tissues are critical to maintain the quiescent state of the vasculature in most adult tissues and organs (1) . Although it is believed that production of these inhibitors is decreased in actively angiogenic tissues, they are often found in tumor tissues or in association with tumor growth, e.g., angiostatin, endostatin, tumstatin, serpin antithrombin, and thrombospondin-1 are present in various tumor tissues (2, 3, 4, 5, 6) . These inhibitors can be produced by tumor cells and/or host cells infiltrated in the tumor (7) . Switching on the expression of these inhibitors at high levels is perhaps one of the host defense mechanisms to prevent tumor growth. In most cases, these angiogenesis inhibitors may suppress tumor growth and keep tumors in a microscopic dormant state by preventing essential neovascularization (8 , 9) . However, most visible tumors switch on their angiogenic phenotypes by overexpression of angiogenic factors, including the families of vascular endothelial growth factor/vascular permeability factor, fibroblast growth factor-2, and angiopoietins (1 , 10, 11, 12, 13, 14) . Almost all angiogenesis inhibitors have been tested or at least considered as therapeutic agents in the treatment of animal cancers. Encouraged by their antitumor effects in animals, many angiogenesis inhibitors, including angiostatin and endostatin, have entered early phases of clinical trials of cancer therapy. Their therapeutic efficacies alone or in combination with other methods in the treatment of human cancers remain to be seen.

Angiostatin, an internal fragment of plasminogen, is a potent and specific endogenous inhibitor of angiogenesis (2) . Despite the fact that it is currently in clinical trials for the treatment of primary tumors and metastasis, the underlying molecular mechanisms and signaling pathways of angiostatin-mediated angiostatic activity remain poorly understood. The structure of angiostatin includes the first three or four kringle domains of plasminogen (2) . Most angiostatic activity of angiostatin is contained within the first three kringles, with kringle 1 (K1) being the most potent single domain (15) . In addition to K1-4, K5 is a potent inhibitor of angiogenesis (16) . A dramatically increased angiostatic activity has been observed in a fragment of plasminogen, called K1-5, containing both angiostatin and most of the K5 domain (17) . Angiostatin and other inhibitors, including thrombospondin-1, endostatin, and tumstatin, are reported to specifically induce endothelial cell apoptosis (18, 19, 20, 21, 22, 23, 24, 25) . However, the role of endothelial apoptosis in antiangiogenic effects of angiostatin and the signaling pathways of angiostatin-induced apoptosis are not well understood.

In addition to endothelial apoptosis, angiostatin binds to an ATP synthase on the surface of human endothelial cells (26 , 27) . However, it is not known if the endothelial cell surface ATP synthase mediates K1-5-induced endothelial apoptosis. Here, we demonstrate that K1-5 binds to the endothelial ATP synthase and that the caspase-mediated endothelial apoptosis is essential for K1-5 to inhibit angiogenesis and tumor growth.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents, Cells, and Animals.
Human plasminogen, K1-3, and K1-5 were prepared and purified to homogeneity according to our methods published previously (17) . An anticaspase-3 monoclonal antibody was obtained as a gift (In Vitro, Stockholm, Sweden). Cell membrane permeable caspase-3 inhibitors, z-DEVD-fmk, z-IETD-fmk, z-LEHD-fmk (Enzyme System Products, Livermore, CA), cell permeable DEVD-CHO (Calbiochem, San Diego, CA), and Ac-DEVD-CHO (Sigma, Stockholm, Sweden) were purchased. The antiendothelial cell surface ATP synthase {alpha}- or ß-subunit antibodies were obtained from Molecular Probes (Eugene, OR). BCE cells were kindly provided by Dr. Judah Folkman (Harvard Medical School, Boston, MA) and maintained as described previously (17) . Rat vascular smooth muscle cells (VSMC) (a kind gift from Dr. Johan Thyberg, Karolinska Institute, Stockholm, Sweden) were grown in 10% heat-inactivated FCS-F12-Ham’s medium and murine T241 fibrosarcoma cells, and Lewis Lung carcinoma cells were grown and assayed in 10% FCS-DMEM. Male 5–6-week-old C57BL/6 mice (Microbiology and Tumor Biology Center, Karolinska Institute) were acclimated and caged in groups of six or fewer. Animals were anesthetized with injections of hypnorm:dormicum:H2O 1:1:2 before all procedures and euthanized with a lethal dose of CO2. All animal studies were reviewed and approved by the animal care and use committee of the Stockholm Animal Board.

Fluorescence-Activated Cell Sorter Analysis.
Proliferating BCE cells were treated with 1 µM K1-5 or plasminogen for 24 h. The cells were trypsinazed and stained with fluorochrome-labeled inhibitors of caspases (FAM-VAD-FMK) according to manufacturer’s protocols (Immunochemistry Technologies, Bloomington, MN). Stained cells were analyzed with a Becton Dickinson FacsScan and CellQuest software.

Detection of Apoptotic Endothelial Cells and Caspase-3 Activation.
Proliferating BCE cells were treated with K1-3, K1-5, or plasminogen (1 µM) in DMEM containing low glucose supplemented with 10% bovine calf serum for 24 h. Cells were harvested and resuspended in PBS with 30 mM glycerol and 0.1 M NaCl, followed by drying on slides and fixation with aceton-methanol (1:1) for 10 min. The fixed cells were stained with ethidium bromide or Hoechst 33258 (500 ng/ml). Apoptotic cells were counted in randomly selected fields using florescent microscope (x20, 10 fields/sample). z-DEVD-fmk, z-IETD-fmk, and z-LEHD-fmk at 20 µM were incubated with the cells for 2 h before addition of K1-5. Apoptotic cells were detected under a fluorescent microscope. BCE cells, treated with cisplatin at 10 µM, K1-3, K1-5, or plasminogen at 1 µM, or various concentrations of K1-5 for 24 h, were lysed with 1% Triton X-100, 20 mM Tris-HCL (pH 8.0), 15 mM NaCl, and 5 mM EDTA. In another experiment, K1-5 at 1 µM was analyzed at different time points. Equal amounts of protein from each sample were separated on 4–12% Bis-Tris gels (Invitrogen, Stockholm, Sweden). Detection of cleaved and activated caspase-3 by a specific monoclonal antibody was performed using the enhanced chemiluminescence plus system (Amersham Biosciences AB, Uppsala, Sweden)

In Vitro Caspase Assays.
Caspase-3-like, caspase-8, and caspase-9 activities were fluorometrically determined by cleavage of cellular substrates of DEVD-7-amino-4-methyl coumarin (DEVD-AMC), IETD-AMC, or LEHD-AMC, respectively, according to a protocol described previously (Peptide Institute, Osaka, Japan; Ref. 28 ). Briefly, ~0.5 x 106 cells were collected at each time point, washed twice in PBS, sedimented by centrifugation, and kept at –20°C. Some samples were preincubated for 1 h with z-DEVD-fmk, z-IETD-fmk, and z-LEHD-fmk (20 µM; Enzyme Systems Products), with 100 mg/ml anti-integrin-{alpha}vß3-neutralizing antibody (Chemicon International, Inc., Temecula, CA) or 100 µg/ml anti-{alpha}- and/or anti-ß-ATPase antibody (Molecular Probes). Frozen cells were resuspended in 25 µl of PBS before measurement and transferred onto a 96-well microtitration plate. The appropriate peptide substrates were mixed in a reaction buffer containing 100 mM 4-morpholinepropanesulfonic acid (pH 6.5; for caspase-9) or 100 mM HEPES (pH 7.0; for caspases-3 and -8), 10% polyethylene glycol, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid, 5 mM DTT, and 10-6% NP40. The cleavages of the fluorogenic peptide substrates were monitored by AMC release in a Fluoroscan II plate reader (Labsystems, Stockholm, Sweden) using {lambda} = 355 nm for excitation and 460 nm for emission. Fluorescence units were converted to picomoles of AMC using a standard curve generated with free AMC. Data were analyzed by linear regression.

Immunofluorescence Analysis.
BCE cells maintained on gelatinized glass coverslips were stained with the anti-ATP synthase antibodies as described previously (27) . Briefly, BCE cells were washed with PBS several times before fixation with 4% paraformaldehyde. A sample was permeabilized with 100% ethanol for 5 min at room temperature as positive control. For immunofluorescent staining, cells were incubated with murine monoclonal antibodies against the {alpha}- or ß-subunit of ATP synthase (Molecular Probes), followed by incubation with a horse antimouse IgG FITC (1:50; Vector Laboratories, Inc., Burlingame, CA). Samples stained only with the secondary antibody served as negative controls. After vigorous washing, cells were mounted and analyzed using a fluorescent microscope with x40 magnification.

ELISA Assay.
Purified recombinant F1 ATP synthase (15 µg/ml) expressed in Escherichia coli bacterial cells was passively adsorbed onto 96-well plates (Immulon 4HBX Flat Bottom Microfilter Plate, VWR International, Batavia, IL). Briefly, plates were coated with recombinant F1 ATP synthase in 50 µl of 0.1 M NaHCO3 (pH 9.6) and incubated overnight at 4°C. Nonspecific sites were blocked with PBS (pH 7.0) containing 1% BSA for 2 h at room temperature. Increasing amounts of K1-5 (0–0.7 M) were added to a final volume of 50 µl. The samples were incubated at 4°C overnight, washed with 0.05% Tween 20 in PBS (pH 7.0), and then incubated with an antihuman angiostatin (goat IgG; R&D Systems, Inc., Minneapolis, MN) antibody for 4 h. After washing, the samples were incubated with biotin-rabbit antigoat IgG (Zymed, San Francisco, CA) for 1 h, followed by incubation with horseradish peroxidase-conjugated streptavidin (Zymed) for 1 h. After a final wash, 100 µl of 3,3',5,5' tetramethylbenzidine (TMB) substrate were added to each well, and the reaction was terminated with 100 µl of 1 M H2SO4. The absorbance of each sample at {lambda} = 450 nm was determined with a Molecular Devices SpectraMax Plus-384 plate reader (n = 4).

Measurement of F1 ATPase Activity.
F1 ATPase activity was measured according to methods described previously (29) . The assay exhibits activity only in the reverse direction, acting to hydrolyze ATP. The hydrolysis of ATP to ADP is coupled to the oxidation of NADH via pyruvate kinase and lactate dehydrogenase. The oxidation of NADH to NAD is read at {lambda} = 340 nm as a decrease in absorbance. Therefore, a decrease in absorbance is a measure of F1 ATPase activity. Briefly, recombinant F1 ATP synthase was coincubated with K1-5 for 30 min before the initiation of the enzymatic reaction. Control studies were performed with azide or in the absence of purified recombinant F1 protein (n = 3).

Chick Embryo Chorioallantoic Membrane (CAM) Assay.
The CAM assay was performed as described previously (17) . Three-day-old fertilized white Leghorn eggs (OVA Production AB, Morgongåva, Sweden) were cracked, and chick embryos with intact yolks were carefully placed in 20 x 100-mm plastic Petri dishes. After 48-h incubation in 4% CO2 at 37°C, disks of methylcellulose containing K1-5 (12.5 µg/mesh) alone, or in combination with the cell permeable DEVD-CHO (1–10 µg/mesh), dried on a nylon mesh (4 x 4 mm) were implanted on the CAM of individual embryos. The nylon mesh disks were made by desiccation of 20 µl of 0.45% methylcellulose (in H2O). After 48–72 h of incubation, embryos and CAMs were examined for the formation of avascular zones in the field of the implanted disks using a stereoscope. Six to nine embryos were used in each group. Values represent mean determinates ± SEM.

Mouse Corneal Micropocket Assay.
The mouse corneal assay was performed according to procedures described previously (17 , 30) . Corneal micropockets were created with a modified von Graefe cataract knife in the eyes of male 6–7-week-old C57BL/6 mice. Micropellets (0.35 x 0.35 mm) of sucrose aluminum sulfate (Bukh Meditec, Copenhagen, Denmark) coated with hydron polymer type NCC containing ~80 ng of fibroblast growth factor-2 alone, a mixture of fibroblast growth factor-2 and Ac-DEVD-CHO (2.5 µg/pellet), Ac-DEVD-CHO alone, or PBS were implanted into the corneal pockets, positioned ~1.2 mm from the corneal limbus. Mice were treated with s.c. injections of either PBS or K1-5 at a dose of 2 mg/kg/day. Corneal neovascularization in each group (n = 10) was examined using a slit-lamp microscope at day 5 after pellet implantation.

Tumor Studies.
Approximately 1 x 106 murine T241 fibrosarcoma tumor cells were implanted s.c. into each C57BL/6 mouse. Six to seven mice were used in the treated and control groups. Intralesional injections with 50 µg of K1-5 alone or in combination with 10 µg of Ac-DEVD-CHO in 100 µl/mouse began shortly after tumor cell implantation and continued once daily for a total of 15 treatments. Visible primary tumors were measured using digital calipers on the days indicated. Tumor volumes were calculated according to the formula: width2 x length x 0.52 as reported previously (17) .

Immunohistochemistry.
For detection of apoptotic endothelial cells in the tumor vasculature, tumor tissues were fixed with 4% paraformaldehyde and embedded using a paraffin method. Sections (5 µm) were dewaxed and rehydrated according to standard protocols. The terminal deoxynucleotidyl transferase-mediated nick end labeling (TUNEL) staining was performed according to a modified fluorescein in situ Death Detection Kit (Amersham). Briefly, the samples, after deparaffinization and rehydration, were blocked using 3% H2O2 in methanol and treated with 20 µg/ml proteinase K (Invitrogen). TUNEL reaction mixture was incubated with samples in a humid atmosphere at 37°C for 1 h. To detect CD31-positive endothelial cells, the sections were blocked with 20% normal rabbit serum and by using avidin-biotin kit (Vector Laboratories). The sections were stained with a rat antimouse monoclonal CD31-biotin antibody at 1:100 (PharMingen, Stockholm, Sweden) and streptavidin conjugated to Cy3 (1:2500; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). The sections were photographed and counted under a fluorescent microscope at x40 magnification.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Induction of Endothelial Apoptosis by K1-5.
We have reported previously that K1-5 potently inhibits BCE cell growth in vitro. To determine whether K1-5 induces endothelial cell apoptosis, purified proteolytically derived human K1-5 at a concentration of 1 µM, known to inhibit endothelial cell growth, was incubated with proliferating BCE cells (17) . After 24-h incubation, a significant proportion of BCE cells became apoptotic as detected by Hoechst 33258 and ethidium bromide staining (Fig. 1, C and GCitation , arrows). The apoptotic BCE cells exhibited characteristic features of cellular apoptosis, including condensation and fragmentation of the nuclei. Proteolytic K1-3 of human plasminogen, known as an active fragment of angiostatin, also induced BCE cell apoptosis under the same conditions (Fig. 1, B and F)Citation . However, apoptotic quantification analysis demonstrated that K1-5 at the concentration of 1 µM was ~5-fold more potent than K1-3 (Fig. 1I)Citation . These data are consistent with our previous finding that K1-5 is a more potent endothelial cell inhibitor than angiostatin. In contrast, intact human plasminogen, the parental molecule of K1-3 and K1-5, did not induce BCE cell apoptosis at the same concentration (Fig. 1, D, H, and I)Citation . As a negative control, BCE cells treated with buffer alone did not exhibit endothelial cell apoptosis (Fig. 1, A, E, and I)Citation . Induction of cellular apoptosis by K1-5 appeared to be endothelial cell specific because vascular smooth muscle and tumor cells were completely insensitive to K1-5 treatments at high concentrations as compared with nontreated cells (Fig. 1, M–P)Citation . The K1-5-induced apoptotic BCE cells were further quantified with fluorescence-activated cell sorter analysis using fluorochrome-labeled inhibitors of caspases as an apoptotic marker, which measures general caspase activity in cells (31, 32, 33, 34) . Again, K1-5 but not the intact plasminogen remarkably induced BCE apoptosis (Fig. 1, J–L)Citation . These findings demonstrate that K1-5 specifically induced endothelial cell apoptosis, which likely involves cascade of caspase activation.



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  		Fig. 1. Induction of endothelial apoptosis. Kringle (K)1-3, K1-5, or plasminogen was incubated with BCE cells for 24 h, followed by staining cells with Hoechst (A–D) or ethidium bromide (E–H). Apoptotic cells were quantified using Hoechst staining and present as means in percentages (±SEM; I). ***, P < 0.001. Fluorescence-activated cell sorter analysis of cell populations with activated caspases of nontreated (J), K1-5-treated (K), and plasminogen-treated (L) BCE cells. Cell morphology stained with Hoechst of K1-5-treated vascular smooth muscle cells (VSMC) (N) and T241 tumor cells (P). Nontreated VSMC (M) and T241 cells (O) were used as controls.

 
Activation of Caspase-3 by K1-5.
The caspase-mediated apo-ptotic pathway plays a central role in regulating cellular growth and death (35) . Among all known caspases, caspase-3 is considered as a main effector of cellular apoptosis. To study whether caspase-3 could be processed and activated by K1-5, highly purified proteolytically derived human K1-5 (1 µM) was added to proliferating BCE cells, followed by detection of the activated form of caspase-3 with a specific antibody. As shown in Fig. 2ACitation , both K1-5 and K1-3 could induce processing of procaspase-3 by converting the precursor molecule into the cleaved active form (arrow). In contrast, human plasminogen was unable to activate caspase-3 (Fig. 2A)Citation . Because K1-5 displayed more potent effects than K1-3 in inducing endothelial apoptosis, we focused our study on K1-5 in subsequent experiments. Additional experiments demonstrated that the activation of caspase-3 by K1-5 was dose dependent (Fig. 2B)Citation . Low levels of activated caspase-3 were detectable at a concentration of 1 nM and plateaued by 1 µM. The maximal activation of caspase-3 occurred after 24-h incubation (Fig. 2C)Citation .



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  		Fig. 2. Activation of caspase-3 and inhibition of kringle (K)1-5-induced endothelial apoptosis and antiangiogenesis by caspase-3 inhibitors. Activation of caspase-3 by K1-3 and K1-5 was detected by a specific antibody against the cleaved form of caspase-3 (arrows in A–C). Activation of caspase-3 by K1-5 was dose- (B) and time- (C) dependent. Proliferating BCE cells were incubated with K1-5 in the presence (E) or absence (D) of z-DEVD-fmk. Apoptotic cells were randomly counted (x20, 10 fields), and percentages of apoptotic cells were presented as means (±SEM; F). ***, P < 0.001. Western blot analysis of activated caspase 3 in the presence and absence of DEVD of K1-5-treated BCE cells (G). Fluorometric analysis of caspase 3 activity of K1-5-treated BCE cells in the presence and absence of DEVD (H). Avascular zones of K1-5-treated CAMs in the presence of DEVD were detected under a microscope (x10) and presented as percentages (n = 5–7; I). *, P < 0.05; **, P < 0.01. K1-5 was administered by s.c. injections (once daily) in mice whose corneas were implanted with fibroblast growth factor (FGF)-2 alone (J) or a combination of FGF-2/DEVD (K). Corneal neovascularization was measured as vessel areas (L; n = 10 corneas). The data were presented as mean determinants (±SEM). ***, P < 0.001.

 
Prevention of Endothelial Apoptosis in Vitro by a Caspase-3 Inhibitor.
If activation of caspase-3 by K1-5 is essential for endothelial cell apoptosis, inhibition of caspase-3 should block K1-5-induced endothelial cell apoptosis. To test this hypothesis, we applied z-DEVD-fmk, as a cell membrane permeable and irreversible caspase-3 inhibitor, to prevent K1-5-induced endothelial cell apoptosis (36 , 37) . At a concentration of 20 µM, z-DEVD-fmk almost completely blocked K1-5-induced BCE cell apoptosis (Fig. 2, D–FCitation ; P < 0.001). These results demonstrate that caspase-3 is an essential and key mediator of K1-5-induced endothelial cell apoptosis. Western blot analysis revealed that DEVD-fmk did not seem to affect processing and activation of caspase-3 induced by K1-5 (Fig. 2G)Citation . However, DEVD-fmk almost completely inhibited caspase-3 activity in K1-5-treated BCE cells (Fig. 2H)Citation .

Impairment of the in Vivo Angiostatic Activity of K1-5 by Caspase Inhibitors.
To demonstrate in vivo that caspase-3-mediated apoptosis is essential for the potent antiangiogenic activity of K1-5, a cell membrane permeable reversible caspase-3 inhibitor, DEVD-CHO, was coimplanted with K1-5 in the chick CAM assay. As expected, K1-5 at the dose of 12.5 µg/mesh induced the formation of avascular zones in all CAMs (n = 7). Interestingly, when K1-5 was combined with DEVD-CHO, the antiangiogenic effect was completely abolished. The opposing antiangiogenic effect was dose dependent and statistically significant (Fig. 2I)Citation .

To further investigate the inhibitory effect of DEVD-CHO on the antiangiogenic activity of K1-5, we performed the mouse corneal angiogenesis assay (17) . The fibroblast growth factor-2-induced corneal angiogenesis was almost completely inhibited by K1-5 treatments at the dose of 2 mg/kg/day (Fig. 2J)Citation . Consistent with the CAM assay, Ac-DEVD-CHO could completely neutralize the antiangiogenic activity of K1-5 in this model (Fig. 2K)Citation . Quantification of corneal neovascularization as vessel areas showed that the antagonistic effect of Ac-DEVD-CHO on the antiangiogenic activity of K1-5 was statistically significant (P < 0.001; Fig. 2LCitation ). These data demonstrate that K1-5-induced antiangiogenesis requires caspase-3-mediated apoptosis.

Neutralization of Antitumor Activity by Caspase Inhibitors Is Correlated with Prevention of Tumor Vasculature from Apoptosis.
We reported previously that administration of K1-5 exhibited potent antitumor activity in a mouse fibrosarcoma model (17) . Consistent with this finding, administration of K1-5 into murine fibrosarcomas at the dose of 50 µg/mouse/day resulted in significant suppression of tumor growth (n = 6; Fig. 3ACitation ). Approximately 83% of the antitumor effect was observed at day 15 after treatment. In contrast, Ac-DEVD-CHO alone (10 µg/mouse/day) did not significantly inhibit tumor growth (Fig. 3A)Citation . The antitumor activity of K1-5 was totally attenuated when K1-5 and Ac-DEVD-CHO were coadministrated into tumors (Fig. 3A)Citation . These results indicate that the antitumor activity of K1-5 requires caspase-3-mediated apoptosis.



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  		Fig. 3. Attenuation of antitumor activity and tumor vascular apoptosis by a caspase-3 inhibitor. In A, kringle (K)1-5 alone or together with Ac-DEVD-CHO was intralesionally injected into murine fibrosarcomas, starting the day of tumor cell implantation and being terminated at day 15. PBS and Ac-DEVD-CHO were used as controls. Tumor volumes were measured every other day according to a standard formula: width2 x length x 0.52. In B, tumor sections (day 15) were double stained with a specific antibody against CD31 (vessels in red) and TUNEL (apoptotic cells in green). Apoptotic microvessels (yellow) were detected by superimposing two images using a digital program. All positive signals were quantified under a microscope (x40, 15 fields), and data were represented as mean determinants (±SEM).

 
To correlate the antitumor effect of K1-5 with its ability to induce microvessel apoptosis, we stained tumor tissues with endothelial- and apoptotic-specific markers. Tumor microvessels were stained with a specific anti-CD31 antibody and labeled with red color (Fig. 3B)Citation . Apoptotic cells in tumor tissues were revealed by TUNEL method labeled with green color. A significant reduction of tumor neovascularization was observed in K1-5-treated tumors (Fig. 3B)Citation . Again, Ac-DEVD-CHO could neutralize this effect as measured by microvessel density. A number of tumor cells were committed to apoptosis as demonstrated by TUNEL staining (Fig. 3B)Citation . Ac-DEVD-CHO only slightly affected overall tumor cell apoptosis in the K1-5-treated tumors as compared with K1-5 alone. A significant number of endothelial cells in tumor vessels underwent apoptosis in the K1-5-treated tumors as revealed by CD31/TUNEL double staining of the same sections (arrows, yellow in Fig. 3BCitation ). Coadministration of Ac-DEVD-CHO with K1-5 significantly prevented endothelial cell apoptosis in microvessels induced by K1-5. These data provide con-vincing evidence that caspase-3 inhibitors prevent K1-5-induced microvessel apoptosis in tumors.

Sequential Activation of Caspases-8, -9, and -3.
To gain additional insights with response to the activation of caspases, we measured the activity of caspases-8 and -9 at various time points. After K1-5 treatment, caspase-8 was highly activated within 3 h. This activation reached a maximal level at ~18 h and persisted for >24 h (Fig. 4A)Citation . Similarly, the maximal activation of caspase-3 was also detected ~18 h after exposure to K1-5 (Fig. 4B)Citation . In contrast, a delayed activation of caspase-9 was observed in K1-5-treated BCE cells (Fig. 4C)Citation . A significant proportion of caspase-9 was activated in ~18 h and reached a maximal level at ~24 h. These results suggest that activation of caspases-8 and -3 precedes caspase-9 activation. To further study the sequential events of caspase activation, we treated BCE cells with various caspase-specific inhibitors. As expected, z-IETD-fmk, a caspase-8-specific inhibitor, efficiently blocked caspases-8 and -9 activities (Fig. 4D)Citation . Similarly, z-LEHD-fmk, a caspase-9-specific inhibitor, completely blocked caspase-9 activity and partially attenuated caspase-8 activation. However, z-DEVD-fmk, a caspase-3 selective inhibitor, blocked caspase-9 activity ~100% and only slighted affected caspase-8 activation. These data suggest that the order of caspase activation in these cells is caspases-8, -3, -9, and -3, again. Like caspase-3 inhibitor, z-IETD-fmk and z-LEHD-fmk were also able to significantly prevent K1-5-induced BCE cell apoptosis (Fig. 4, E–H)Citation .



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  		Fig. 4. Sequential activation of caspases-8, -9, and -3 by kringle (K)1-5. Activation of caspases-8 (A), -3 (B), and -9 (C) by K1-5 at different time points was measured by cleavage of specific substrates. At 24 h after incubation of K1-5, activation of caspases-8 and -9 was measured in the presence of caspase-3 inhibitor, z-DEVD-fmk; caspase-8 inhibitor, z-IETD-fmk; and caspase-9 inhibitor, z-LEHD-fmk (D). BCE cell morphological changes were examined at 24 h after incubation with K1-5 in the absence (E) or presence of z-IETD-fmk (F) or z-LEHD-fmk (G). Apoptotic cells were quantified by randomly counting apoptotic bodies in 10 fields (x20). **, P < 0.01; ***, P < 0.001.

 
K1-5 Binds to Endothelial Cell Surface ATP Synthase and Triggers Caspase Activation.
Although K1-5 was able to induce endothelial cell apoptosis by activation of intracellular caspases, this finding could not explain the specific inhibitory activity of K1-5 on endothelial but not other cells. To identify endothelial cell surface-specific molecules that could potentially mediate K1-5-induced apoptosis, we tested the possibility that K1-5 could bind to endothelial cell surface ATP synthase, identified as an angiostatin-binding molecule specifically expressed on endothelial cells (26 , 27) . We first demonstrated that BCE cells used in our system expressed both the {alpha}- and ß-subunits of ATP synthase (Fig. 5, B and C)Citation . This interaction was specific because FITC-labeled antibody alone did not result in positive staining (Fig. 5A)Citation . A sensitive ELISA analysis demonstrated that K1-5 bound to purified, recombinant ATP synthase in a dose-dependent manner (Fig. 5D)Citation . We further demonstrated that K1-5 significantly inhibited F1 ATP synthase reverse activity (Fig. 5E)Citation . Having shown that K1-5 bound to endothelial cell surface ATP synthase, we performed studies to correlate this interaction with K1-5-induced endothelial apoptosis. Two specific neutralizing antibodies against the {alpha}- and ß-subunits of ATP synthase were added to endothelial cells in the presence and absence of K1-5. Interestingly, both anti-ATP synthase antibodies sufficiently blocked K1-5-induced caspases-3, -8, and -9 activities (Fig. 5, F–H)Citation . Previously, it has been reported that angiostatin binds to endothelial cell surface integrin-{alpha}vß3 (38) . To test whether integrin-{alpha}vß3 could play a role in mediation of K1-5-induced caspase activation and endothelial cell apoptosis, a neutralizing antibody against integrin-{alpha}vß3 was incubated with endothelial cells in the presence of K1-5. It appeared that the anti-integrin-{alpha}vß3-neutralizing antibody had only a little effect (<17%) on K1-5-induced caspase-3 activity, whereas it had no effect on K1-5-induced endothelial cell apoptosis (Fig. 5, I and J)Citation . These data demonstrate that interference of ATP synthase activation with antibodies blocks K1-5-induced endothelial cell apoptosis. Thus, endothelial cell surface ATP synthase plays a critical role in K1-5-induced caspase activation and endothelial cell apoptosis.



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  		Fig. 5. Kringle (K)1-5 binding to endothelial cell surface ATP synthase. BCE cells were stained with an anti-ATP synthase {alpha}-subunit antibody (B), anti-ATP synthase ß-subunit antibody (C), or without a primary antibody (A). A sandwich ELISA assay detected the binding of the F1 subunit of ATP synthase to K1-5 in a dose-dependent manner (D). Reverse ATP synthase activity was determined in the presence and absence of K1-5 (E). Blockage of activation of caspases-8 (F), -3 (G), and -9 (H) induced by K1-5 by the anti-{alpha} or -ß ATP synthase subunit antibodies. Cell apoptosis (I) and caspase activity (J) in the presence of an anti-integrin {alpha}vß3-neutralizing antibody of K1-5-treated BCE cells.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, we provide compelling evidence that K1-5-induced endothelial cell death contributes to its antiangiogenic mechanism. Since the discovery of angiostatin as the first endothelial cell-specific inhibitor produced in association with tumor growth (2) , great efforts have been devoted to understanding the underlying mechanisms and signaling pathways. Despite these efforts, how this molecule specifically acts on endothelial cells remains an enigma. Although many interesting hypotheses have been raised to explain the specific effect of angiostatin on growing endothelial cells, these theories generally require additional experimental support. As both angiostatin and K1-5 are relatively large protein molecules, which cannot easily penetrate the cell membrane, it would be expected that the endothelial cell target molecules for angiostatin and K1-5 would be located on the cell surface. Therefore, several laboratories have focused on identifying endothelial cell surface molecules that bind to or interact with angiostatin. One intriguing study demonstrates that angiostatin, but not plasminogen, binds to ATP synthase on the surface of endothelial cells with its catalytic subunits located extracellulary (26) . It appears that inhibiting this enzyme with a specific antibody impairs the antiproliferative effect of angiostatin (26) . More recently, it was shown that angiostatin inhibits ATP synthase activity on endothelial cells (27) . Another group used a genetic approach to identify an angiostatin-binding protein in endothelial cells (39) . They have found that recombinant angiostatin expressed as a cytosolic protein interacts with angiomotin, an endothelial cell protein involved in motility (39) . In addition to endothelial cell surface ATP synthase and angiomotin, integrin {alpha}vß3 has been reported to bind to angiostatin (38) . However, it has been difficult to study whether angiomotin or integrin {alpha}vß3 transduces apoptotic signals triggered by angiostatin due to lack of specific neutralizing antibodies against bovine capillary endothelial angiomotin and other caspase components. Thus, it is important that further work should be devoted to develop these valuable reagents to know if these components other than endothelial cell surface ATP synthase also contribute to angiostatin/K1-5-induced endothelial apoptosis. However, our present findings suggest that it is unlikely that integrin {alpha}vß3 plays significant roles in mediating K1-5-induced endothelial apoptosis. Furthermore, our present work explains, at least in part, the underlying mechanisms by which angiostatin/K1-5 specifically inhibits endothelial cell growth. If K1-5 binds to endothelial cell surface ATP synthase, it explains why K1-5 specifically acts on endothelial cells but not other cell types, because this endothelial cell surface molecule is only present on endothelial cells. Our findings that specific inhibition of various caspase activities completely impairs the antiproliferative, antiangiogenic, and antitumor effects of K1-5 suggest that caspase-mediated cellular apoptosis is essential for the angiostatic function of K1-5. Can we link K1-5-induced, caspase-mediated endothelial apoptotic pathways to its interaction with endothelial cell surface ATP synthase? The fact that antibodies against the {alpha}- and ß-subunits of ATP synthase, known to block angiostatin-induced endothelial proliferation, can block K1-5-induced caspase activities clearly demonstrates that this cell surface ATP synthase is involved in triggering the intracellular caspase pathway. Our study on the cascade event of caspase activation indicates sequential activation of caspases-8, -3, and -9 (shown in schematic Fig. 6Citation ). Although the molecular details of these events deserve additional studies, it is clear that caspase-8 becomes activated first. The activation of caspase-8 by endothelial cell surface ATP synthase remains to be investigated. In addition to angiostatin and K1-5, several other endogenous angiogenesis, including endostatin, tumstatin, and thrombospondin-1, have been reported to induce caspase activation and endothelial cell apoptosis (18 , 21, 22, 23, 24, 25) . The relation between their endothelial cell surface receptors and induction of caspase activity/endothelial cell apoptosis remains to be further studied.



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  		Fig. 6. Schematic representation of the role of kringle K1-5-induced endothelial apoptotic pathways in regulation of angiogenesis and tumor growth. K1-5 binds to endothelial cell surface ATP synthase and sequentially activates caspases-8, -3, -9, and -3 again that lead to endothelial cell apoptosis. The apoptotic commitment of endothelial cells results in its specific suppression of angiogenesis and tumor growth.

 
Angiostatin, endostatin, and several endogenous angiogenesis inhibitors, either alone or in combination with other therapeutic methods, have produced remarkable efficacy in suppression of primary and metastatic tumor growth in animal models (40, 41, 42) . These promising animal studies have led to clinical trials of these inhibitors in suppression of human tumors. At the time of submitting this study, it still remains to be seen if these angiogenesis inhibitors are potent tumor suppressors in humans. Our present study provides important insight into the molecular mechanisms for understanding how these angiogenesis inhibitors act on newly formed blood vessels. Although several questions remain to be addressed, including the mechanisms behind angiogenesis inhibitors’ differential effect on active and quiescent endothelial cells and the roles of angiogenesis inhibitors in physiological and pathological angiogenesis, we believe that our work provides important clues for further clinical development of these angiogenesis inhibitors in cancer therapy.


   ACKNOWLEDGMENTS
 
We thank Hong Xu and Aimee Paradis for technical help and advice.


   FOOTNOTES
 
Grant support: Human Frontier Science Program, the Swedish Cancer Foundation, the Karolinska Institute Foundation, the Åke Wibergs’ Foundation, the Swedish Research Council, the Swedish Heart and Lung Foundation, and the National Cancer Institute (CA-86344). Y. Cao is supported by the Karolinska Institute and Swedish Research Council.

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.

Requests for reprints: Yihai Cao, Microbiology and Tumor Biology Center, Karolinska Institutet, S-171 77 Stockholm, Sweden. Phone: (46)-8-728 7596; Fax: (46)-8-31 94 70; E-mail: yihai.cao{at}mtc.ki.se

Received 6/16/03. Revised 3/ 6/04. Accepted 3/11/04.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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