Structural elements of kallistatin required for inhibition of angiogenesis

Robert Q. Miao, Vincent Chen, Lee Chao, and Julie Chao

Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina 29425 - 2211


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Kallistatin is a serpin first identified as a specific inhibitor of tissue kallikrein. Our recent studies showed that kallikrein promoted angiogenesis, whereas kallistatin inhibited angiogenesis and tumor growth. This study is aimed to identify the structural elements of kallistatin essential for its antiangiogenic function. Kallistatin mutants at the hinge region (A377T) and a major heparin-binding domain (K312A/K313A) were created by site-directed mutagenesis. Recombinant kallistatin mutant A377T did not bind or inhibit tissue kallikrein activity. Wild-type kallistatin and kallistatin mutant A377T, but not kallistatin mutant K312A/K313A lacking heparin-binding activity, inhibited VEGF-induced proliferation, growth, and migration of human microvascular endothelial cells. Similarly, wild-type kallistatin and kallistatin mutant A337T, but not kallistatin mutant K312A/K313A, significantly inhibited VEGF-induced capillary tube formation of cultured endothelial cells in Matrigel and capillary formation in Matrigel implants in mice. To elucidate the role of the heparin-binding domain in modulating angiogenesis, we showed that wild-type kallistatin interrupted the binding of 125I-labeled VEGF to endothelial cells, whereas kallistatin mutant K312A/K313A did not interfere with VEGF binding. Consequently, wild-type kallistatin, but not kallistatin mutant K312A/K313A, suppressed VEGF-induced phosphorylation of Akt. Taken together, these results indicate that the heparin-binding domain, but not the reactive site loop of kallistatin, is essential for inhibiting VEGF-induced angiogenesis.

tissue kallikrein; heparin-binding domain; reactive site loop; serpin


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

ANGIOGENESIS IS THE FORMATION of new capillaries from preexisting blood vessels and is a multistep process involving endothelial cell proliferation, migration, and capillary tube formation via differentiation (12, 26). Under physiological conditions, angiogenesis is a highly regulated phenomenon and is under the control of angiogenic stimulators and inhibitors (4). However, unregulated angiogenesis happens in ischemia-dependent diseases, such as tumor growth and diabetic retinopathy (5, 17). Under ischemic conditions, hypoxia activates a hypoxia-inducible factor-binding sequence in the VEGF promoter. This leads to transcription of VEGF mRNA and increased production of VEGF protein. VEGF mRNA is substantially upregulated in most human tumors (18). There is a correlation between VEGF expression and microvessel density in primary breast cancer sections (20). These findings suggest that ischemia-induced VEGF expression tips the balance between negative and positive angiogenesis regulators, and VEGF plays an important role in ischemia-induced angiogenesis.

Serine proteinase inhibitors (serpins) have coevolved with serine proteinases and are the principal regulators of a variety of divergent serine proteinases (35). Inhibitory serpins interact with their target proteinases via a reactive site loop structure of 30-40 amino acids from the COOH terminus (35). Serpins participate in a variety of physiological events, such as coagulation, fibrinolysis, complement activation, and phagocytosis, by targeting serine proteinases (35). However, serpins have alternative functions independent of their regulation of proteolytic events, such as hormone transport and cell differentiation (35). Recently, several serpins, including pigment epithelium-derived factor (PEDF) (13), antithrombin (32), and maspin (40), have been shown to have antiangiogenic activity. The amount of inhibitory PEDF produced by retinal cells is directly correlated with oxygen concentrations. Hyperoxia induces the expression of PEDF, whereas hypoxia decreases the PEDF levels in retinal cells (13). Therefore, downregulation of PEDF and upregulation of VEGF during hypoxic conditions facilitate neovascularization (13, 21). PEDF as a naturally occurring angiogenesis inhibitor plays a crucial role in maintaining the angiogenesis balance. Like PEDF, endogenous kallistatin levels markedly decrease in vitreous fluid of patients with diabetic retinopathy compared with those of healthy individuals (30). Therefore, kallistatin may be involved in modulating angiogenesis-based diabetic retinopathy.

Kallistatin was first identified as a tissue kallikrein-binding protein (7, 8). Tissue kallikrein belongs to a subset of closely related serine proteinases that exhibit a narrow range of substrate specificity (3). Tissue kallikrein is involved in the processing of kininogen to produce biologically active peptide kinins. A recent study (15) showed that the expression of the genes encoding bradykinin B1 and B2 receptors and tissue kallikrein was increased in skeletal muscle of ischemic hindlimbs. Kallikrein gene delivery promoted angiogenesis in ischemic hindlimbs, and ischemia-induced spontaneous angiogenesis was abolished in the bradykinin B1 receptor knockout mice or by chronic infusion of B1 receptor antagonist (16). Conversely, native kallistatin has been shown to inhibit VEGF- and basic fibroblast growth factor (bFGF)-induced proliferation, migration, and adhesion of cultured endothelial cells and microvessel formation in Matrigel implants in mice (31). Kallistatin gene delivery suppresses spontaneous angiogenesis in the ischemic hindlimb and tumor growth in nude mice (31). Whether the inhibitory effect of kallistatin on ischemia-induced angiogenesis is dependent on its inhibitory activity toward tissue kallikrein remains unclear. We have created a kallistatin mutant (A377T) lacking the inhibitory activity toward tissue kallikrein and a kallistatin mutant (K312A/K313A) lacking the heparin-binding activity by using site-directed mutagenesis. The effects of recombinant wild-type kallistatin and kallistatin variants on the VEGF-induced angiogenesis were evaluated in vitro and in vivo to identify the essential structural elements of kallistatin required for regulating angiogenesis.


    MATERIALS AND METHODS
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INTRODUCTION
MATERIALS AND METHODS
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Cell culture. Human dermal microvascular endothelial cells (HMVEC) (Clonetics, San Diego, CA) were cultured in EGM-2-MV medium (Clonetics) supplied with 5% FBS; 10 ng/ml human EGF, bFGF, and VEGF; and 50 µg/ml gentamicin. Experiments were performed between cell passages 4 and 10.

Site-directed mutagenesis of kallistatin variants. Construction, expression, purification, and characterization of recombinant kallistatin were performed as previously described (9). The strategy of site-directed mutagenesis for kallistatin mutant A377T was based on a previously described method (28). All the mutant fragments were synthesized by PCR by using oligonucleotides containing the desired mutations in the primers. For A377T, two separate PCR fragments were generated and treated with appropriate restriction enzymes for each variant. One fragment, primed by oligonucleotides 5'-GGCACAATCCAGCTTATC-3' and 5'-CGGTGCCAGCCTCATCC-3' and digested by XhoI, together with the other fragment, primed by oligonucleotides 5'-AGGAAGCAGCAGCCACCAGCTT-3' and 5'-CTATGGTTTCGTGGGGT-3' with a mutant codon (underlined) and digested by SalI, were ligated into XhoI/SalI-digested pTrc-HKBP to generate A377T. Kallistatin mutant K312A/K313A was created by double mutation of two lysine residues in the major heparin-binding domain as previously described (11).

Analysis of complex formation by SDS-PAGE. The ability of native and recombinant kallistatins to form SDS-stable complexes with tissue kallikrein was assessed by incubating 2 µM of different kallistatins with 1 µM tissue kallikrein in 50 mM Tris · HCl, pH 8.0, and 0.1 M NaCl, at 37°C for certain time periods as follows: plasma and wild-type recombinant kallistatin for 10 min and kallistatin mutant A377T for 4 h. Reactions were quenched by adding SDS sample buffer containing DTT boiled for 2 min and analyzed by SDS-PAGE on a 10% polyacrylamide gel.

Determination of association rate by kinetic assays. Association rate constants of the recombinant kallistatins and human tissue kallikrein were determined under pseudo-first-order conditions as previously described (10). Inhibitory reactions of recombinant kallistatins toward human tissue kallikrein were performed at 37°C in 1 ml of reaction buffer containing 20 mM sodium phosphate, pH 8.0, 100 mM NaCl, and 0.1% BSA. An excess of inhibitors, wild-type and variant kallistatins, were incubated with 3 nM human tissue kallikrein at various enzyme-to-inhibitor molar ratios. At certain incubation intervals, 100 µl of the reaction mixture were removed and added immediately to 30 µM of D-Val-Leu-Arg-MCA in 2 ml of buffer containing 50 mM Tris · HCl buffer, pH 8.0. Residual enzymatic activity of tissue kallikrein was measured at 25°C by monitoring the release of 7-amino-4-methylcoumarin at the wavelength of 380-nm excitation and 460-nm emission in a PerkinElmer LS-50 luminescence spectrometer. Residual enzymatic activity was proportional to the initial velocity obtained through the slope of the absorbance plotted against time. The apparent rate constant, kobs, was calculated by using the equation ln([Et]/[E0]) -kobs t. [E0] represents the initial enzymatic activity of tissue kallikrein without the addition of kallistatin. [Et] represents the remaining enzymatic activities of tissue kallikrein at different intervals of incubation with kallistatin. The association rate constant (kass) was calculated by using the equation kass= kobs/[I], where [I] represents the concentration of the inhibitor in the assay mixture and kobs is the apparent rate constant.

[3H]thymidine incorporation. Cell proliferation assay was performed by [3H]thymidine incorporation as previously described (43). Quiescent HMVEC in 48-well plates were treated with 15 ng/ml VEGF (VEGF165 was kindly provided by Genentech, South San Francisco, CA) without or with different concentrations of kallistatin in fresh 2% FBS medium 199 for 18 h and then pulse labeled with 1 µCi/ml of [3H]thymidine (DuPont New England Nuclear, Boston, MA) for another 6 h. Radioactivity was determined by using a liquid scintillation counter (Packard, Downers Grove, IL).

Cell growth assay. Cell growth assay was performed as previously described (39). In brief, HMVEC were plated at a concentration of 2 × 104 cells/well in 24-well plates and allowed to attach for 24 h in endothelial cell basal medium (Clonetics) supplemented with 0.5% FBS, and then was treated with VEGF of 15 ng/ml with or without kallistatin. After 3 days incubation, cell number was determined by using a hemacytometer after staining with 0.2% trypan blue.

Cell migration assay. Cell migration was assessed by using a modified Boyden chamber (Costar transwell inserts; Corning, Acton, MA) (42). The transwell inserts were coated with a solution of 100 µg/ml type I collagen (Sigma, St. Louis, MO) at 4°C overnight and then air dried. VEGF (15 ng/ml), with or without kallistatin (1 µM) dissolved in endothelial cell basal medium containing 0.1% BSA, was added in the bottom chamber of the Boyden apparatus. HMVEC (2 × 105 cells) suspended in a 100-µl aliquot of endothelial cell basal medium containing 0.1% BSA were added to the upper chamber. After 4 h of incubation, cells on both sides of the membrane were fixed and stained with a Diff-Quik staining kit (Dade Division, Baxter Healthcare, Miami, FL). The average number of cells from five randomly chosen high-power (×400) fields on the lower side of the membrane was counted.

Competitive binding of 125I-labeled VEGF with kallistatin to endothelial cells. Competitive binding of 125I-labeled VEGF with kallistatin to the endothelial cell surface was performed as previously described (27). Quiescent HMVEC were incubated with 1 ng/ml 125I-labeled VEGF in binding buffer (DMEM supplemented with 20 mM HEPES, pH 7.4, and 0.1% BSA) at 4°C for 2 h in the absence or presence of increasing concentrations of kallistatin or kallistatin mutants. Unbound 125I-labeled VEGF was removed by washing three times with cold binding buffer and once with cold PBS (pH 7.4). Bound protein was solubilized in 250 µl of 0.3 M NaOH, and radioactivity was determined by using a Multigamma counter (Pharmacia, Turku, Finland). Nonspecific binding was determined by measuring binding in the presence of a 100-fold molar excess of unlabeled VEGF. Specific binding was determined by subtracting the nonspecific binding from the total binding.

Western blot analysis of Akt phosphorylation. Total cell lysates were prepared by adding 200 µl of cell lysate buffer containing (in mM) 20 Tris · HCl, pH 7.5, 150 NaCl, 1 EDTA, 1 EGTA, 2.5 sodium pyrophosphate, 1 Na3VO4, 1 phenylmethylsulfonyl fluoride, and 1% Triton X-100, and 1 µg/ml leupeptin as previously described (33). Total cell extract (60 µg) was separated on a 7.5-15% gradient SDS-PAGE gel and transferred to a Hybond enhanced chemiluminescence nitrocellulose membrane (Amersham, Piscataway, NJ). Phosphorylation of Akt kinase and total Akt levels were determined by using a rabbit polyclonal phosphospecific Akt antibody (1:1,000 dilution; New England BioLabs, Beverly, MA) and rabbit polyclonal Akt antibody (1:1,000 dilution; New England BioLabs) as previously described (29).

Capillary tube formation assay in vitro. Capillary tube formation assay on Matrigel was performed as previously described (29). Matrigel (Becton Dickinson, Franklin Lakes, NJ) was used to coat a 24-well plate at 4°C and allowed to polymerize at 37°C for 30 min. HMVEC were seeded (5 × 104 cells/well) on Matrigel-coated plates. Cells were incubated with VEGF (15 ng/ml) with or without kallistatin or kallistatin mutants (1 µM) in endothelial cell basal medium containing 2% FBS. After cells were incubated for 18 h, capillary tube formation was examined visually under a phase-contrast microscope. The intact tube number in six random views of × 100 magnification was counted.

Matrigel angiogenesis model in mice. Male FVB mice (6-8 wk; Harlan Sprague Dawley, Indianapolis, IN) were subcutaneously injected with 0.5 ml Matrigel (9-10 mg/ml) containing 150 ng/ml VEGF, with or without 2 µM kallistatin or kallistatin mutants, near the abdominal midline by using a 26-gauge needle as previously described (34). One week after Matrigel injection, mice were killed, and the Matrigel plug, along with overlying skin and peritoneal membrane, was removed and fixed in 4% buffered formaldehyde in PBS. Plugs were embedded in paraffin, sectioned, and stained by using Masson's trichrome staining for histological analysis. The cell-occupied area per field of view (3.46 × 104 µm2) from 10-15 fields (×400 magnification) in the tissue sections was measured by using a computerized digital camera system and NIH Image 1.61. The vessels are defined as those structures possessing a patent lumen and positive endothelial nucleus. The study complied with the standards for care and use of animal subjects as stated in the Guide for the Care and Use of Laboratory Animals [DHEW Publication No. (NIH) 85-23, Revised 1985, Office of Science and Health Reports, DRR/NIH, Bethesda, MD 20205]. All animal procedures were approved by the Animal Care and Use Committee of the Medical University of South Carolina.

Statistical analysis. Statistical significance was determined by one-way ANOVA with the Fisher multiple comparison test. All data are expressed as means ± SE, and differences were considered significant at a value of P < 0.05.


    RESULTS
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ABSTRACT
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RESULTS
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Expression and purification of recombinant wild-type kallistatin and kallistatin mutants. The positions for site-directed mutations of human kallistatin at the hinge region and heparin-binding domain are shown in Fig. 1A. The mutation site of A377T is located at P12 [by using the nomenclature by which the scissile bond occurs between residues designated P1 (F388) and P1' (S389)] in the hinge region of the reactive center loop. The double mutation sites of K312A/K313A are located at the major heparin-binding domain between C2 sheet and H helix (11). The constructs were verified by DNA sequencing. Recombinant kallistatins were expressed in an Escherichia coli expression system as an intracellular protein under the direction of the Trc promoter. The wild-type and kallistatin variants in cell extracts were identified as a single band of 46 kDa by Western blotting with the use of a monoclonal antibody (G4C10) against kallistatin (9).


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Fig. 1.   A: positions for site-directed mutagenesis of kallistatin. The residue of alanine (P12) at the hinge region was substituted with threonine to create kallistatin variant A377T lacking the inhibitory activity toward tissue kallikrein. The two residences of lysine at the major heparin domain between C2 sheet and H helix were substituted with alanine to create kallistatin mutant lacking heparin-binding activity. Mutant sites are bold and underlined. B: complex formation of purified recombinant kallistatins with tissue kallikrein. Reactions of various recombinant kallistatins (2 µM) with human tissue kallikrein (1 µM), carried out at 37°C, were analyzed by SDS-PAGE. Lane 1, molecular mass standard; lane 2, human tissue kallikrein; lane 3, native kallistatin alone; lane 4, native kallistatin incubated with human tissue kallikrein for 10 min; lane 5, recombinant wild-type kallistatin alone; lane 6, recombinant wild-type kallistatin incubated with human tissue kallikrein for 10 min; lane 7, A377T alone; and lane 8, A377T incubated with human tissue kallikrein for 4 h.

Analysis of kallistatin and tissue kallikrein complex formation by SDS-PAGE. Inhibition of tissue kallikrein by kallistatin was accompanied by formation of a SDS- and heat-stable high molecular weight complex of 85 kDa. Figure 1B shows Coomassie blue staining of purified recombinant kallistatins and kallistatin-kallikrein complexes in SDS-PAGE. Purified recombinant wild-type kallistatin and kallistatin mutant migrated as a single band of ~46 kDa on SDS-PAGE. A lack of glycosylation in the prokaryotic expression system resulted in recombinant kallistatin having a lower molecular mass compared with the 58 kDa plasma kallistatin. The complex formation was visualized after 10-min incubation of tissue kallikrein with plasma kallistatin (lane 4) or recombinant wild-type kallistatin (lane 6). The mutant A377T, containing a mutation at the hinge region, did not form a complex even after 4-h incubation with tissue kallikrein; however, it was cleaved by human tissue kallikrein (lane 8). These results indicate that replacement of alanine at P12 with threonine impairs the activity of kallistatin to form a stable complex with tissue kallikrein. Moreover, this substitution at the hinge region converts kallistatin from an inhibitor to a substrate of tissue kallikrein.

Inhibitory activities of wild-type kallistatin and kallistatin mutants toward tissue kallikrein. Kinetic analysis of the inhibition of human tissue kallikrein by recombinant kallistatins was performed under pseudo-first-order conditions by using an excess of recombinant kallistatins at various molar ratios to tissue kallikrein. For native and recombinant kallistatins, the concentrations at 0.15, 0.2, 0.25, 0.3, and 0.35 µM were used. The recombinant kallistatins inhibited the activity of tissue kallikrein in a concentration- and time-dependent (0 to 15 min) manner (9). The kallistatin variant A377T containing mutation at P12 at the hinge region had much lower inhibitory activities toward tissue kallikrein than the wild-type kallistatin. To observe the inhibition of kallikrein by this variant, the molar ratio of kallistatin/kallikrein was increased to 500 and 1,000 in the reaction mixture. The kass is 1.42 × 104 M-1 · s-1 for native kallistatin and 1.64 × 104 M-1 · s-1 for recombinant kallistatin (Table 1). The kass of A377T variant was estimated to be <10 M-1 · s-1. In contrast, kallistatin mutant K312A/K313A maintained its inhibitory activity toward tissue kallikrein and its kass is 0.8 × 104 M-1 · s-1. The association rate constants of native kallistatin and recombinant wild-type kallistatin were similar, indicating that the addition of a hexahistidine sequence at the amino terminus did not affect the inhibitory activity of the recombinant kallistatin. These results support the fact that short side-chain residues in the hinge region are required for kallistatin to adopt an inhibitory conformation.

                              
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Table 1.   kass values for inhibition of human tissue kallikrein by kallistatins

Effect of wild-type kallistatin and kallistatin mutants on the proliferation, growth, and migration of cultured endothelial cells. Purified recombinant wild-type kallistatin and kallistatin mutants were used to determine their effect on the proliferation, growth, and migration of cultured human endothelial cells. Cell proliferation was determined by [3H]thymidine incorporation. Exogenous recombinant wild-type kallistatin and kallistatin mutant A377T lacking inhibitory activity toward tissue kallikrein dose-dependently inhibited VEGF-induced endothelial cell proliferation compared with the VEGF control (Fig. 2, A and B). However, contrary to wild-type kallistatin, kallistatin mutant K312A/K313A lacking heparin-binding activity had no effect on VEGF-induced endothelial cell proliferation (Fig. 2C). Similarly, both recombinant wild-type kallistatin and kallistatin mutant A377T, but not kallistatin mutant K312A/K313A, dose-dependently suppressed VEGF-induced endothelial cell growth measured by cell counts (Fig. 3). Furthermore, the effect on VEGF-induced endothelial cell migration was also determined by using modified Boyden chambers. Similarly, recombinant wild-type kallistatin and kallistatin mutant A377T, but not kallistatin mutant K312A/K313A, also significantly inhibited VEGF-induced endothelial cell migration compared with the VEGF control (Fig. 4). The effect of kallistatin on endothelial cell migration is not as potent as its effect on cell proliferation and growth. These results indicate that the heparin-binding domain of kallistatin is essential for its inhibition of VEGF-induced endothelial cell proliferation and migration, whereas the kallikrein inhibitory activity is not required for this inhibition.


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Fig. 2.   Effect of recombinant wild-type kallistatin and kallistatin mutants on VEGF-induced proliferation of human dermal microvascular endothelial cells (HMVEC). Quiescent HMVEC were incubated with the indicated concentrations of recombinant wild-type human kallistatin (HKBP) or kallistatin mutants (A377T and K312A/K313A) and VEGF (15 ng/ml) for 24 h. DNA synthesis was measured as [3H]thymidine incorporation. Results are expressed relative to controls without any treatments. Each value represents the mean ± SE (n = 4).



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Fig. 3.   Effect of recombinant wild-type kallistatin and kallistatin mutants on VEGF-induced growth of HMVEC. Quiescent HMVEC were incubated with the indicated concentrations of recombinant wild-type HKBP or kallistatin mutants (A377T and K312A/K313A) and VEGF (15 ng/ml) for 72 h. Cell growth was determined by cell counting. Each value represents the mean ± SE (n = 4).



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Fig. 4.   Effect of recombinant wild-type kallistatin and kallistatin mutants on VEGF-induced migration of HMVEC. Cell migration was determined by using modified Boyden chambers. VEGF: 15 ng/ml; recombinant wild-type HKBP or kallistatin mutants (A377T and K312A/K313A): 1 µM and 0.1 µM. The average number of cells from five randomly chosen high power (× 400) fields on the lower side of the membrane was counted. Each value represents the mean ± SE (n = 3).

Effect of recombinant kallistatin and kallistatin mutants on VEGF-induced capillary tube formation of cultured endothelial cells in Matrigel in vitro. Capillary tube formation assay in Matrigel is a useful in vitro assay to determine the branching morphogenesis of endothelial cells, which is a complex developmental program that regulates the formation of the blood vessels. Figure 5A shows representative images of endothelial cell tube formation in Matrigel. VEGF significantly induced tube formation compared with the control (21 ± 1 vs. 4 ± 1 tubes/field, P < 0.001, n = 3) (Fig. 5B). However, recombinant wild-type kallistatin (HKBP) and noninhibitory variant of kallistatin (A377T) has an identical inhibitory effect on VEGF-induced capillary tube formation in Matrigel (10 ± 0.5 or 11 ± 0.5 vs. 21 ± 1 tubes/field, P < 0.001, n = 3). Conversely, kallistatin mutant K312A/K313A lacking heparin-binding activity lost the capability to inhibit VEGF-induced capillary tube formation of endothelial cells in Matrigel in vitro (20 ± 0.5 vs. 21 ± 1 tubes/field, P < 0.001, n = 3).


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Fig. 5.   Effect of recombinant wild-type kallistatin and kallistatin mutants on VEGF-induced capillary tube formation of endothelial cells on Matrigel. A: representative images of tube formation on Matrigel. HMVEC were treated with recombinant wild-type kallistatin (1 µM) and kallistatin mutants (A377T and K312A/K313A at 1 µM) in the presence of VEGF (15 ng/ml) for 18 h. B: quantitation of tube number on Matrigel. Intact tube number was counted in 6 random view at the magnification of ×100. Each value represents the mean area ± SE (n = 3).

Effect of wild-type kallistatin and kallistatin mutants on VEGF-induced angiogenesis in subcutaneously implanted Matrigel in mice. Based on the in vitro studies, we used the animal model of subcutaneously implanted Matrigel to evaluate the antiangiogenic effect of kallistatin and its variants in vivo. Figure 6A shows representative images of Matrigel implants stained with Masson's trichrome. VEGF significantly increased capillary density compared with the control. Red blood cells appeared abundantly in the capillaries, indicating that these capillaries are functional. However, the mean capillary area in the Matrigel implant containing both VEGF and kallistatin (HKBP) was significantly lower than that in the VEGF group (1,418 ± 104 vs. 4,707 ± 471 µm2/field, n = 5-6, P <0.001) (Fig. 6B). Noninhibitory variant of kallistatin (A377T) has the identical inhibitory effect on VEGF-induced angiogenesis in Matrigel implant as the recombinant wild-type kallistatin (1,231 ± 146 vs. 4,707 ± 471 µm2/field, n = 5-6, P <0.001). However, kallistatin variant (K312A/K313A) lacking heparin-binding activity lost its ability to inhibit VEGF-induced angiogenesis in vivo (5,221 ± 592 vs. 4,707 ± 471 µm2/field, n = 5, P = 0.283). These data indicate that the ability of kallistatin to suppress VEGF-induced angiogenesis in Matrigel implants in mice is dependent on its heparin-binding domain but independent of its inhibitory activity toward tissue kallikrein.


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Fig. 6.   Effect of recombinant wild-type kallistatin and kallistatin mutants on VEGF-induced angiogenesis in Matrigel implants. A: histological analysis of implanted Matrigel plugs. Matrigel mixed without or with VEGF (150 ng/ml) in the presence or absence of 2 µM kallistatin (HKBP) was injected subcutaneously into mice. The Matrigel plugs were stained with Masson's trichrome at 1 wk after injection. B: quantitation of mean capillary area in implanted Matrigel plugs. Each value represents the area mean ± SE (n = 6).

Effect of wild-type kallistatin and kallistatin mutant K312A/K313A on 125I-labeled VEGF binding to cultured endothelial cells and phosphorylation of Akt kinase in cultured endothelial cells. Kallistatin is a heparin-binding protein. The major heparin-binding domain of kallistatin has been localized in the region between C2 sheet and H helix (11). Double mutation of two lysine residues to alanine (K312A/K313A) resulted in marked reduction of heparin-binding activity (11). Kallistatin may act by competing with VEGF binding to heparan-sulfate proteoglycans, a low affinity-binding site, and thus suppressing VEGF-binding activity and the angiogenesis signaling cascades induced by VEGF. To test this hypothesis, we examined the capacity of kallistatin to inhibit VEGF binding to endothelial cells. Figure 7A demonstrates that recombinant wild-type kallistatin dose-dependently inhibits VEGF binding to HMVEC. In the presence of 1 µM of recombinant wild-type kallistatin, the specific binding of 125I-labeled VEGF was reduced by 40% (Fig. 7A). However, recombinant kallistatin mutant K312A/K313A lacking heparin-binding activity did not interfere with VEGF binding to endothelial cells (Fig. 7A). Nonspecific binding (2,532.6 ± 112.9 cpm/well) is 21.5% of VEGF total binding to human endothelial cells (11,783.4 ± 516.9 cpm/well). The effect of kallistatin on VEGF-induced Akt phosphorylation in cultured endothelial cells was further determined by Western blot analysis. Results show that wild-type kallistatin, but not kallistatin mutant K312A/K313A lacking heparin-binding activity, abolished VEGF-induced phosphorylation of Akt kinase in cultured endothelial cells (Fig. 7, B and C). These results further support those in vitro and in vivo findings that heparin-binding domain is involved in inhibiting VEGF-induced angiogenesis.


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Fig. 7.   Effect of kallistatin on the specific binding of 125I-labeled VEGF to HMVEC and VEGF-induced phosphorylation of Akt kinase in HMVEC. A: kallistatin inhibits the specific binding of 125I-labeled VEGF to HMVEC. Increasing concentration of recombinant wild-type kallistatin or kallistatin mutant (K312A/K313A) at the heparin binding domain was incubated with HMVEC in the presence of 125I-labeled VEGF (1 ng/ml) for 2 h at 4°C. Nonspecific binding was determined in the presence of a 100-fold molar excess of unlabeled VEGF and was subtracted from total binding. Results were represented as the percentage of VEGF binding in the absence of competitors (noncompetitive binding). Each value represents the mean ± SE (n = 4). B: kallistatin suppresses VEGF-induced phosphorylation of Akt kinase in HMVEC. Quiescent HMVEC were incubated without any treatment (control) or with 15 ng/ml VEGF in the presence or absence of 1 µM wild-type kallistatin or kallistatin mutant K312A/K313A for 15 min. Cell lysate (60 µg) was used in Western blot analysis of phosphorylated Akt kinase and total Akt kinase. C: quantitative results of Western blot analysis of phosphorylated Akt kinase and total Akt kinase (n = 3).


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The hinge region in the reactive-center loop of a serpin is important in maintaining its inhibitory activities toward serine proteinases. The insertion of the reactive-center loop into the beta -sheet A is required for a serpin to adopt a stable inhibitory conformation (6). The sequence of the hinge region is mostly composed of short-chain amino acids, such as alanine and glycine, which are the most conserved residues in the reactive site loop. These short-chain residues in the hinge region confer reactive-center loop flexibility and thus allow the insertion of the loop. Replacement of small amino acids with bulky amino acids in the hinge region of kallistatin may hinder this insertion and thereby decelerate the conformational change required for inhibitory activity. Our results showed that replacement of alanine residue with threonine at the P12 position at the hinge region converts kallistatin variant A377T from inhibitor to substrate for tissue kallikrein. Similarly, a single mutation from alanine to proline within the reactive center loop of other inhibitory serpins results in conversion of the serpin from a proteinase inhibitor to a proteinase substrate (22). These findings support the hypothesis that a partial insertion of the hinge region results in a constrained loop that could adopt a canonical form for the recognition by a serine proteinase (6). Taken together, our results showed that the short-chain amino acid residues in the hinge region of kallistatin are important to maintain the inhibitory conformation required for inhibiting tissue kallikrein activity.

In this study, we showed that like wild-type kallistatin, kallistatin mutant A377T lacking the inhibitory activity toward human tissue kallikrein, still exerted the same inhibitory effect on VEGF-induced angiogenesis in vitro and in vivo. These results indicate that the reactive site loop of kallistatin is crucial to its ability to inhibit tissue kallikrein but not essential to inhibit angiogenesis. Several serpins have been shown to have antiangiogenesis activities independent of their ability to inhibit proteinases. For example, PEDF is a noninhibitory serpin (2). Antithrombin (32) and maspin (40), after cleavage of their reactive site loop in the COOH terminus, still maintain their potent antiangiogenic activity. It also has been reported that several serpins, such as PEDF, antithrombin, and plasminogen activator inhibitor (PAI-1), bind to components of the extracellular matrix (1). The ability of PAI-1 to inhibit angiogenesis in vivo is partially attributed to its binding to vitronectin, not its proteinase inhibitory activity (38).

VEGF is a key growth factor stimulating angiogenesis and triggers the phosphatidylinositol 3-kinase-Akt signal transduction pathway, which appears to mediate many cellular responses associated with the regulation of angiogenesis (25). VEGF is a heparin-binding growth factor, and binding of VEGF to heparan-sulfate proteoglycans on endothelial cell surfaces can enhance the binding of VEGF to VEGF receptor (Flk-1/KDR) (23). Kallistatin is also a heparin-binding protein (41). The major heparin-binding domain of kallistatin was identified in the region between the H helix and C2 sheet of kallistatin that contains clusters of positively charged residues (11). Site-directed mutagenesis of the basic residues within this domain (K312A/K313A) resulted in a loss of specific heparin-binding capacity (11). Our in vitro studies showed that recombinant wild-type kallistatin, but not kallistatin mutant K312A/K313A, suppressed the VEGF-induced proliferation, growth, migration, and capillary tube formation of endothelial cells in Matrigel. Therefore, we speculate that kallistatin may compete with VEGF binding to heparan-sulfate proteoglycans, a low affinity-binding site, and thus suppress VEGF binding activity and VEGF-induced signaling required for angiogenesis. In support of this hypothesis, we performed the competitive binding assay of 125I-labeled VEGF with wild-type kallistatin and kallistatin mutant K312A/K313A. Results showed that recombinant wild-type kallistatin, but not kallistatin mutant K312A/K313A lacking heparin-binding activity, competed with the specific binding of 125I-labeled VEGF to endothelial cells and further suppressed the VEGF-induced phosphorylation of Akt in endothelial cells. These results suggest that kallistatin interferes with VEGF binding at the heparin-binding site and the heparin-binding domain in kallistatin may play an important role in the inhibition of angiogenesis. This action of kallistatin is similar to that of endostatin as the inhibitory effect of endostatin on bFGF-induced angiogenesis is mediated by its heparin-binding activity (14, 36). In many receptor systems, ligand binds first to an abundant low-affinity receptor, which draws the ligand onto the cell surface and then transfers it to a second, high-affinity receptor that transduces the appropriate signal into cells (19). The most common and widely acting, low-affinity receptors are the heparan sulfate proteoglycans, which play central roles in the reception and modulation of a wide range of growth factors. Heparan sulfate proteoglycans also act in combination with membrane integrins to control cell adhesion and migration (19). In this regard, kallistatin is expected to be a broad-spectrum inhibitor capable of inhibiting angiogenesis mediated by VEGF, bFGF, and other heparan sulfate-dependent growth factors.

The inhibitory effect of kallistatin on angiogenesis was further demonstrated by an in vivo Matrigel implantation model. Matrigel is a laminin-rich reconstituted matrix extracted from the Englebreth-Holm-Swarm tumor. This model has proved suitable for assessing angiogenesis and antiangiogenic agents (24, 34, 37). Matrigel not only can trap the growth factor to allow slow release and prolonged exposure to surrounding tissues but can also enhance the selectivity of endothelial cells entering the gel plug, because basement membranes are not readily crossed by fibroblasts and certain other cells (34). Previous studies showed that results of angiogenesis assay using the in vivo Matrigel model are consistent with those using the classic rat cornea angiogenesis assay (24). Our results show that VEGF significantly induced angiogenesis and high density of capillaries with red blood cells in Matrigel plug containing VEGF alone. However, the capillary area is markedly smaller and capillary density is significantly lower in wild-type kallistatin or kallistatin mutant A377T-treated Matrigel plug than those in Matrigel plug containing VEGF alone. In contrast, the heparin-binding domain mutant of kallistatin (K312A/K313A) lost its inhibitory activity against VEGF-induced angiogenesis in the implanted Matrigel in mice, indicating that the heparin-binding domain of kallistatin, not its reactive site loop, determines its inhibitory effect on VEGF-induced angiogenesis. Although this study showed that the inhibitory effect on VEGF-induced angiogenesis was not attributed to its proteinase inhibitory activity, kallistatin may be involved in the inhibition of kinin-induced angiogenesis by inhibiting tissue kallikrein activity in vivo. This hypothesis needs to be examined in a future study by delivering kallistatin mutants in the ischemic hindlimb model in which kinin has been shown to promote the blood flow recovery (15).

In conclusion, kallistatin markedly inhibits angiogenesis by suppressing VEGF-induced proliferation, growth, and migration of endothelial cells. The heparin-binding domain, not the reactive site loop of kallistatin, is essential for its inhibitory activity. The results clearly demonstrate that kallistatin acts like a noninhibitory serpin in inhibiting VEGF-induced angiogenesis.


    ACKNOWLEDGEMENTS

This work was supported by the National Heart, Lung, and Blood Institute Grant HL-44083.


    FOOTNOTES

Address for reprint requests and other correspondence: J. Chao, Dept. of Biochemistry and Molecular Biology, Medical Univ. of South Carolina, Charleston, SC 29425-2211 (E-mail: chaoj{at}musc.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

10.1152/ajpgi.00524.2002

Received 12 November 2002; accepted in final form 12 February 2003.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Am J Physiol Cell Physiol 284(6):C1604-C1613
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