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Article |
Genetic alteration of endothelial heparan sulfate selectively inhibits tumor angiogenesis
Correspondence to Jeffrey D. Esko: jesko{at}ucsd.edu
To examine the role of endothelial heparan sulfate during angiogenesis, we generated mice bearing an endothelial-targeted deletion in the biosynthetic enzyme N-acetylglucosamine N-deacetylase/N-sulfotransferase 1 (Ndst1). Physiological angiogenesis during cutaneous wound repair was unaffected, as was growth and reproductive capacity of the mice. In contrast, pathological angiogenesis in experimental tumors was altered, resulting in smaller tumors and reduced microvascular density and branching. To simulate the angiogenic environment of the tumor, endothelial cells were isolated and propagated in vitro with proangiogenic growth factors. Binding of FGF-2 and VEGF164 to cells and to purified heparan sulfate was dramatically reduced. Mutant endothelial cells also exhibited altered sprouting responses to FGF-2 and VEGF164, reduced Erk phosphorylation, and an increase in apoptosis in branching assays. Corresponding changes in growth factor binding to tumor endothelium and apoptosis were also observed in vivo. These findings demonstrate a cell-autonomous effect of heparan sulfate on endothelial cell growth in the context of tumor angiogenesis.
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Introduction |
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Binding of both FGF-2 and VEGF164 depends on the sulfation state of heparan sulfate, in particular N-sulfation of glucosamine residues in specific domains along the heparan sulfate chain (Yayon et al., 1991; Turnbull et al., 1992; Ono et al., 1999). A family of four N-acetylglucosamine N-deacetylase/N-sulfotransferases (Ndst) catalyze this reaction (Esko and Selleck, 2002), but microvascular endothelial cells only express Ndst1 and Ndst2 (Wang et al., 2005). Ndst2-deficient mice are essentially normal, aside from defective granule formation in connective tissue-type mast cells (Forsberg et al., 1999; Humphries et al., 1999). In contrast, mice with a systemic deletion of Ndst1 die perinatally because of forebrain defects, skeletal malformation, and lung hypoplasia (Ringvall et al., 2000; Grobe et al., 2005; Pan et al., 2006; Pallerla et al., 2007), but vasculogenesis and angiogenesis up to birth appear normal by gross examination and general histology. Here, we show that altering Ndst1 expression in endothelial cells alters growth of syngeneic murine tumors because of a unique alteration in the tumor microvasculature. In contrast, the mutation had no effect on physiological angiogenesis, e.g., during cutaneous wound repair. The change in tumor angiogenesis appears to result from altered binding and signaling by the proangiogenic growth factors FGF-2 and VEGF.
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Results |
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Physiological angiogenesis during cutaneous wound repair also appeared normal. After a biopsy punch wound of dorsal skin, an intense angiogenic response was detected 3–4 d after injury at the granulating edges of healing wounds. Both mutant and wild-type mice showed a similar response in clustering of CD31-positive processes at the wound margin (Fig. 1 a). No significant difference in the density of processes at the wound edges was noted between the two groups (Fig. 1 b). Vascular density fell with distance from the wound margin in a similar fashion in both mutant and wild-type mice as well. The rate of wound contraction over the 4-d period also did not differ (Fig. 1 c), and in separate experiments, complete resolution of wounds occurred in both groups of mice by 10 d. Analysis of retinal vascular development under normoxic conditions showed only a very minor effect that was not significant (Friedlander, M., personal communication). These findings suggest that physiological angiogenesis was unaffected by endothelial inactivation of Ndst1.
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Tumor microvasculature in mutant mice is altered
To determine whether the decrease in tumor mass was caused by a change in tumor microvasculature, we examined the pattern of staining of CD31 (PECAM-1 [platelet/endothelial cell adhesion molecule-1]) in tumor sections (Fig. 3).
Microvascular density was significantly reduced in tumors grown on the mutant background (Fig. 3 b; 26.8 ± 4.7 vs. 35.3 ± 4.7 microvessels per 400x field in mutant and wild-type mice, respectively; P < 0.02). A slight increase in vessel caliber was also apparent when antibody binding was visualized by fluorescence microscopy (Fig. 3 b).
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Altered binding of FGF-2 and VEGF164 to endothelial heparan sulfate
To gain insight into the mechanism underlying the alteration in tumor vascularization, we attempted to purify endothelial cells from tumors and normal tissues. However, we were unable to obtain sufficient numbers of cells from tumors, and the cultures were overtaken rapidly by small numbers of contaminating tumor cells. In contrast, highly purified cells were obtained from lungs and livers of mutant and wild-type mice. These cells proliferated for two to three passages in culture, and each population maintained their differentiated phenotype based on CD31 expression (Fig. 4 a), positive staining for von Willebrand Factor, and uptake of DiI-labeled acetylated low-density lipoprotein (Wang et al., 2005).
Both mutant and wild-type endothelial cells grew at comparable rates in culture (in medium supplemented with heparin, endothelial cell growth supplement, and 20% FBS; Fig. 4 b). Omission of heparin from the growth medium mildly decreased growth of both mutant and wild-type cells. Thus, isolated cells showed no obvious effect of Ndst1 deficiency on proliferation in monolayer culture.
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Altered signaling responses of mutant endothelial cells to FGF-2 and VEGF164
Under standard culture conditions, wild-type and mutant primary endothelia proliferated at similar rates and to the same extent (Fig. 4 b), but striking differential growth responses of cells were observed when cells were cultivated on Matrigel and supplemented with FGF-2 or VEGF164. Wild-type endothelia proliferated and formed extensive branched networks in response to 1–10 ng/ml FGF-2, whereas mutant cells were unresponsive (Fig. 5, a and b).
Increasing the concentration of FGF-2 to values in excess of the physiological range (100 ng/ml) was sufficient to stimulate a sprouting response by mutant cells (Fig. 5 b, right), indicating that they expressed functional FGF receptors that were not activated at lower doses of FGF-2. Treating wild-type cells with heparinase, which degrades heparan sulfate, mimicked the phenotype of the mutant (Fig. 5 c). The addition of unfractionated heparin also blocked the response, whereas it had no effect on mutant cells (even at low concentration). Mutant cells also failed to respond to VEGF164, whereas wild-type cells sprouted robustly in response to this factor (Fig. 5 d). Infecting endothelial cells derived from Ndst1f/fTekCre– mice with Adenoviral-Cre yielded comparable results (unpublished data), indicating that the effect was cell autonomous and not the result of selection of unresponsive cells during their cultivation.
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Because the Ndst1 deficiency affected vessel-associated VEGF–VEGFR-2 complexes (Fig. 7 b), we examined whether tumor growth depended on VEGFR-2 expression. When LLC tumors were grown in VEGFR-2 heterozygous mice, mean tumor volumes were reduced by 
50% compared with the wild type (P < 0.0001; Fig. 7 f). CD31-stained tumor sections from VEGFR-2 mutants also exhibited reduced microvascular density (33% reduction relative to wild type; P < 0.03) and increased vessel caliber (Fig. 7 g) similar to that described for tumors grown in Ndst1f/fTekCre+ mice (Fig. 3).
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Discussion |
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Recently, Jakobsson et al. (2006) showed that VEGF signaling in endothelial cells and angiogenesis were fully supported by heparan sulfate expressed in trans by adjacent perivascular smooth muscle cells. Given the selective deletion of Ndst1 in endothelial cells in Ndst1f/fTekCre+ mice and the fact that pericyte density was unaltered (Fig. 7 d), it would appear that pericyte heparan sulfate will not substitute for endothelial heparan sulfate during tumor angiogenesis. Thus, in the context of tumor vasculature, endothelial heparan sulfate behaves in a cell-autonomous manner. Ideally one would like to further potentiate the phenotype by more dramatic changes in heparan sulfate composition or content. However, complete ablation of sulfation (Ndst1–/–Ndst2–/–) or inhibition of the polymerase that catalyzes formation of the heparan sulfate chains (Ext1/Ext2) result in early developmental defects, including failure to differentiate mesodermal precursors of the endothelium (Lin et al., 2000; Holmborn et al., 2004; Stickens et al., 2005), thus preventing further studies of the vasculature in adult animals.
Vasculogenesis (endothelial cell differentiation/de novo vessel formation) and physiological angiogenesis appear to be unaffected in Ndst1f/fTekCre+ mice based on normal birth weight, growth of the animals, reproductive capacity, and normal cutaneous wound healing (Fig. 1). Nevertheless, endothelial cells derived from the lungs of mutant animals showed alterations in growth factor binding, signaling, and sprouting behavior under conditions that promote proliferation in vitro (Figs. 4–6
). Why, then, was angiogenesis selectively altered in the tumor environment but not under conditions of cutaneous wound repair? One possibility is that the tumor microenvironment places an especially high demand on the tumor vascular bed with respect to transfer of nutrients, removal of waste products, and gas exchange. If correct, then further stress, e.g., by hypoxia or induced tissue regeneration, might reveal underlying differences in angiogenic capacity in normal tissues. Alternatively, the composition of growth factors might differ in the wound environment, or other glycosaminoglycans, such as dermatan sulfate, which is abundant in wound tissue, might compensate for reduced heparan sulfate in the endothelium (Penc et al., 1998; Taylor et al., 2005). Further studies are needed to address this point.
As observed in other tumor models, the microvasculature in LLC tumors displayed a highly chaotic morphology compared with normal vasculature (Carmeliet and Jain, 2000; Jain, 2005). Altering heparan sulfate specifically in the endothelium resulted in less branched and tortuous tumor vessels and decreased overall density of the neovasculature (Fig. 3). Similar morphological changes have been observed in tumor microvasculature after in vivo treatment with anti-VEGF antibodies or VEGF receptor tyrosine kinase inhibitors (Yuan et al., 1996; Jain, 2003), suggesting that altering heparan sulfate in the endothelium may preferentially affect VEGF-mediated responses. Other observations that support this idea include the following: (1) transfection of VEGF nullizygous tumor cells with VEGF164 rescues tumor growth and vascular density/branching, whereas altered tumor growth and a hypobranched/hypodense vasculature results from transfection with the non–heparin binding isoform, VEGF120 (Grunstein et al., 2000); (2) mouse embryos engineered to express only VEGF120 exhibit a marked decrease in capillary branching accompanied by increased vascular caliber (Ruhrberg, 2003); (3) altering Ndst1 expression reduces the formation of VEGF–VEGFR complexes in mutant tumor vasculature (Fig. 7 b); and (4) reduction of expression of VEGFR2 in heterozygous mutants resulted in reduced tumor growth and vascularization (Fig. 7, f and g). The modest dilatation of mutant tumor microvasculature that we observed could reflect incomplete inactivation of VEGF signaling, as the deficiency in Ndst1, although nearly fully penetrant, does not completely reduce sulfation of the heparan sulfate chains (Fig. 4 e).
The potentiation of growth factor action by heparan sulfate proteoglycans is thought to depend on formation of ternary complexes in which the heparan sulfate chain acts as a scaffold to approximate ligand and receptor, thus affecting signaling intensity and duration (Forsten-Williams et al., 2005). In the extracellular matrix, heparan sulfate proteoglycans also act as reservoir for growth factors (Vlodavsky et al., 1990), providing stability and enhancing the capacity for gradient formation (Lander et al., 2002). Many tumors secrete heparanase, which can degrade the extracellular matrix and liberate bound growth factors (Elkin et al., 2001; Vlodavsky et al., 2002), and overexpression of heparanase in tumor cells results in enhanced angiogenesis and tumor growth (Zcharia et al., 2004). Because endothelial cells produce matrix proteoglycans, loss of Ndst1 activity could, in theory, result in reduced storage of growth factors in the surrounding matrix. Thus, reduced tumor growth in Ndst1f/fTekCre+ mice could reflect a combination of cell-autonomous growth factor signaling defects as well as reduced availability of growth factors in the matrix.
Heparan sulfate chains do not exist as free polysaccharides but, rather, occur covalently bound to proteoglycan core proteins. The major heparan sulfate proteoglycans in endothelial cells consist of the glycosylphosphatidylinositol-linked glypicans, membrane-spanning syndecans, and secreted proteoglycans (perlecan, agrin, and collagen XVIII). Some of these have roles in angiogenesis based on mutational studies. Mutants defective in syndecan-1 show an increase in vessel length during corneal angiogenesis, possibly because of differences in release of inflammatory mediators (Gotte et al., 2002), whereas overexpression of syndecan-1 causes abnormal angiogenesis during cutaneous wound repair (Elenius et al., 2004). In contrast, syndecan-4 deficiency impairs fetal vessels in the placental labyrinth (Ishiguro et al., 2000) and angiogenesis during wound repair (Echtermeyer et al., 2001). Deletion of exon 3 of perlecan core protein, which bears a major heparan sulfate containing domain, impairs wound angiogenesis as well as tumor angiogenesis (Zhou et al., 2004). Mice lacking collagen XVIII show delayed regression of blood vessels postnatally in the vitreous along the surface of the retina and abnormal outgrowth of retinal vessels (Fukai et al., 2002) but enhanced intimal neovascularization in atherosclerotic aortas (Moulton et al., 2004). The variation in phenotype in mutants and tissues may reflect differences in the expression of individual proteoglycans in diverse vascular beds, other functional domains within the core proteins (O'Reilly et al., 1997; Bix and Iozzo, 2005), or changes in heparan sulfate in the supporting tissue in systemic mutants rather than cell-autonomous effects on endothelial cells. Additional studies are needed to analyze the effect of altering specific proteoglycans selectively in endothelial cells.
Our findings validate heparan sulfate and the enzymes involved in its biosynthesis as potential targets for chemotherapeutic intervention. Targeting heparan sulfate could prove advantageous over strategies that focus on individual proangiogenic growth factors, such as VEGF (e.g., bevacizumab; Bergsland, 2004), as heparan sulfate can modulate the activity of multiple angiogenic factors (e.g., hepatocyte growth factor, PDGF, heparin-binding EGF, angiopoietin, tumor necrosis factor-
, interleukin-8, and others; Blackhall et al., 2001; Jiang and Couchman, 2003), most of which bind to heparan sulfate. Previous studies have shown that tumor cell proliferation also depends on heparan sulfate (for review see Fuster and Esko, 2005). Because heparan sulfate biosynthesis in endothelial cells and tumor cells utilizes a common set of enzymes, agents that target heparan sulfate would provide a two-prong approach for blocking both tumor cell division and formation of the supporting vasculature.
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Animals
C57Bl/6 mice harboring a conditional mutation in exon 2 of Ndst1 (Ndst1f/f) were crossed with transgenic mice expressing Cre recombinase under control of the Tek (Tie2 receptor) promoter/enhancer (TekCre+; Kisanuki et al., 2001; Wang et al., 2005). Ndst1f/f TekCre+ males were mated with Ndst1f/f TekCre– females generating equal numbers of mutant and wild-type animals. Genotyping of mice was performed by PCR using genomic DNA from tail biopsies as described previously (Wang et al., 2005). Mice heterozygous for VEGFR-2 (VEGFR-2+/–) were purchased from The Jackson Laboratory, with genotyping performed using primers as described previously (Shalaby et al., 1995). All animals were fully backcrossed on C57Bl/6 background (>10 generations). Mice were housed under barrier conditions in AAALAC-approved vivaria, following standards and procedures approved by the local Institutional Animal Care and Use Committee. They were weaned at 3 wk, maintained on a 12-h light–dark cycle, and fed water and standard rodent chow ad libitum.
Quantitative PCR
To assess the efficiency of Ndst1 inactivation, quantitative real-time PCR (Applied Biosystems) on genomic DNA was performed using primers flanking the downstream loxP recombination site of the Ndst1 floxed construct (Grobe et al., 2005; n = 3 per group). Cycle threshold values from duplicate assays were used to determine the initial starting quantity of loxP DNA from mutant versus wild-type genomic DNA samples.
To examine endothelial Ndst1-4 mRNA, total RNA was isolated from mutant versus wild-type primary lung endothelia, reverse transcribed (Superscript III; Invitrogen), and amplified using gene-specific intron-spanning primers (primer sequences: Ndst1 forward, GGACATCTGGTCTAAG, and reverse, GATGCCTTTGTGATAG; Ndst2 forward, TGGTCCAAGGAGAAAACCTG, and reverse, GGTACGACCTCCGAGTCAAA; Ndst3 forward, CCACTGCCTTGTGTC, and reverse, GGAGTACGCTCGGTC; Ndst4 forward, CTAACTACTTCCACTC, and reverse, ATGTGCACTGCATACC). Cycle threshold values from triplicate assays were used to calculate fold expression compared with expression of ß-actin in the same sample.
Tumor growth studies
Tumor cells were harvested with Trypsin/EDTA (Invitrogen) and were inoculated subcutaneously at 4 x 105 cells per mouse in 50 µl DME into the left hindquarters of 10–12 wk-old mice. Tumor growth was measured using calipers over a 15-d period, and tumor volume was estimated using the following formula: vol = length x (width)2/2. Mice were killed after 15 d (beyond which tumor ulceration became frequent), and their tumors were dissected, measured directly with calipers, and removed for cryo- and formalin fixation for further histologic analysis. Mouse bone marrow transplantation studies were performed as previously described (Wang et al., 2005). In a pilot experiment, all nontransplanted irradiated mice died within 1 wk. Conversely, all mice that underwent marrow transplantation survived until termination of the tumor experiment without ill appearance.
Immunohistochemical analysis of tumor microvasculature
Formalin-fixed and deparaffinized embedded sections from 15-d subcutaneous tumors were stained with hematoxylin/eosin. Labeling of tumor vasculature was performed using rat anti–mouse CD31 (1:200; BD Biosciences) and biotinylated goat anti–rat secondary antibody (1:100; BD Biosciences) and detected using HRP-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories). Development with AEC chromogen (Vector Laboratories) was followed by counterstaining with Mayer's hematoxylin. Microvascular density on tumor sections was quantified by examining the number of vessels per 400x microscopic field over five fields per section. Fluorescence staining was performed using biotinylated secondary antibodies and treatment with Cy3-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories) on frozen tumor sections. In a separate experiment, sections were incubated with rabbit anti-vWF antibody (DakoCytomation), and bound antibodies were detected using HRP-conjugated anti–rabbit antibody (1:100; Jackson ImmunoResearch Laboratories) followed by AEC chromogen/Mayer's counterstain. Microvasculature was quantified by a pathologist blinded to slide identification. Staining of tumor leukocytes and smooth muscle actin was accomplished on paraffin-embedded sections exposed to biotin-conjugated rat anti–mouse CD45 antibody (1:200; BD Biosciences) or rabbit anti–mouse SmA antibody (1:50; NeoMarkers), respectively, with secondary antibodies, development, and counterstaining as described above. Density of SmA-positive processes was quantified for each tumor section by calculating the mean number of stained vessels per 400x field among six fields per section. Fluorescence-stained tumor microvasculature was examined in cryopreserved tumor specimens by acetone fixation of frozen sections, exposure to anti-CD31 mAb, biotinylated secondary antibodies, and treatment with Cy3-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories). Vascular endothelial apoptosis in tumor sections was measured on acetone-fixed frozen sections treated with anti-CD31 mAb, biotinylated secondary antibodies, and AP-conjugated streptavidin followed by development with Vector Blue AP Substrate kit (Vector Laboratories) according to the manufacturer's instructions. Colabeling with TUNEL was performed on the same sections (In Situ Cell Death Detection kit; Roche) and developed using AEC chromogen (Vector Laboratories) according to the manufacturer's instructions.
In situ microvascular FGF-2 reactivity was assessed on frozen tissue sections by incubating specimens with biotinylated FGF-2 (Bai et al., 1999), and bound factor was detected with HRP-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories). For detection of VEGF–VEGFR2 complexes, acetone-fixed frozen sections were labeled with biotinylated GV39M antibody (1:50; East Coast Biologics) followed by treatment with Cy3-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories). Laminin in the same sections was visualized by treatment with rabbit anti-laminin antibody (1:500; DakoCytomation) followed by FITC-labeled goat anti–rabbit antibody (1:50; Chemicon).
All microscopy for immunohistology was performed using a microscope (Axiolab; Carl Zeiss MicroImaging, Inc.) with 20x (Plan-Neofluar/0.5), 40x (Achroplan/0.65), or 100x oil (Plan-Neofluar/1.30) objectives, with all image acquisition taken at room temperature. A 3CCD camera (DKC 5000; Sony) was used for digital image acquisition and captured using NIH ImageJ (Scion Image v. 1.60) software. Image processing and storage (TIFF format) was performed using Photoshop software (Adobe). All fluorescence images were captured using epifluorescence (source for excitation was FluoArc [Carl Zeiss MicroImaging, Inc.]) with appropriate barrier filters on the same microscope system and using fluorochromes applied to tissues/cells as described.
Intravital studies of tumor angiogenesis
LLC LL/2 cells were resuspended in 10 ml complete medium at (2 x 106 cells/ml) and were rocked in a 50-ml silicon-coated flask on a gyratory shaker in culture. After 48–72 h, tumor spheroids of similar diameters (600–1,000 µm) were selected, washed in serum-free medium, and individually placed into dorsal skinfold chambers using a sterile plastic pipette as previously described (Borgstrom et al., 1996). Observations of tumor angiogenesis were made day 12 after tumor spheroid implantation in mutant and wild-type mice as described previously (Manegold et al., 2003). Unanesthetized mice were placed in a plexiglass tube containing a longitudinal slit from which the chamber projected for exposure to the microscope. The tube was immobilized on a plexiglass platform, which was placed on the stage of an intravital microscope (Biomed; Leitz) for viewing the tumor spheroid microcirculation. Images for all mice were recorded at room temperature through a silicone intensified tube camera (SIT68; DAGE-MTI) attached to the microscope (Nikon) using 4x or 10x water-immersion objectives. The camera was connected to a monitor (Panasonic) and an S-VHS video recorder (HC-6600; JVC). For each chamber, the tumor spheroid vasculature was identified using a 4x objective, and regions of highest tumor vascular density were recorded with the 10x objective. Still images of four to six tumor spheroid fields per chamber were recorded at 100x magnification and scanned for grid intersection using a 12 x 12 grid overlay that overlies the image, and branch points per field were measured using Photoshop CS2. Digitized images were also analyzed on Volocity 2.6.1 software (Improvision), which was used to obtain total linear (skeletal) length of vasculature per field, as well as mean vascular caliber per field.
Mouse wound induction and contraction
Under general anesthesia, mice were wounded using a 3-mm-diameter full-thickness dorsal skin punch biopsy. Wounds were then photographed daily over 4 d after injury, and wound area was measured using ImageJ software. The wounds were biopsied to include the margin to normal surrounding skin and snap-frozen in Tissue-Tek OCT (Sakura), and median sections were taken to include the wound fibrin plug at center, flanked by the wound granulation tissue and surrounding normal dermis/epidermis. Acetone-fixed sections were incubated with biotinylated anti–mouse CD31 (1:100; BD Biosciences), followed by HRP-conjugated streptavidin (1:500; Jackson ImmunoResearch Laboratories). Wound microvascular density for any given mouse was then determined by calculating the mean number of CD31+ processes per 400x field using four fields surrounding the wound center.
FGF-2 and VEGF164 binding to mutant endothelial cells and heparan sulfate
Primary lung endothelial cells were harvested at the first or second passage using 5 mM EDTA in PBS. The cells were treated with heparin (1 mg/ml in PBS), washed with PBS, and incubated with biotinylated 0.6 µg/ml FGF-2 in buffer (0.5% BSA/2 mM EDTA in PBS) at 4°C for 1 h with gentle stirring on an orbital shaker (Bai et al., 1999). Bound FGF-2 was detected by flow cytometry using streptavidin-phycoerythrin. A set of cells was exposed only to streptavidin-phycoerythrin as a negative control. To detect VEGF binding, cells (4 x 106/ml) were incubated for 1 h at 4°C with recombinant human biotinylated VEGF165 (1.1 µg/ml in PBS; R&D Systems). Bound VEGF165 was detected with avidin-FITC reagent (1:1,000; R&D Systems) and flow cytometry. As controls, some cells were incubated with an irrelevant biotinylated protein (soybean trypsin inhibitor) or Alexa Fluor 488–conjugated EGF (Invitrogen; each at 8 µg/ml for 45 min at 4°C) followed by flow cytometry.
Primary lung endothelia were labeled for 48 h with 37 µCi/ml H235SO4 (655 Ci/mmol; NEN Life Science Products) in reduced-sulfate medium (Ham's F12 containing 10% dialyzed, filter-sterilized FBS). Isolation of purified heparan [35S]sulfate from cultured cells was performed essentially as described previously (Bame and Esko, 1989). Binding of 0.2 µg FGF-2 and 1 µg VEGF164 was measured using a nitrocellulose filter binding assay using 5,000 cpm per assay (Kreuger et al., 2003). Controls included competition with 100 µg/ml heparin or use of bovine serum albumin in place of growth factor.
[3H]Heparan sulfate was prepared from primary endothelial cells by culturing them in growth medium containing [6-3H]glucosamine and 1 mM reduced glucose. Radiolabeled chains were degraded with heparinases and analyzed by anion-exchange HPLC as described previously (Toyoda et al., 1997).
Angiogenesis assays on reconstituted extracellular matrix
Murine lung endothelial cells were isolated from 8–12-wk-old mutant mice or their wild-type littermates as described previously (Wang et al., 2005). Purity was assessed by flow cytometry using CD31 mAb (FACSCalibur; BD Biosciences), uptake of DiI-acetylated-LDL (10 µg/ml; Biomedical Technologies), and reaction with polyclonal antibody to von Willebrand factor (DakoCytomation).
Matrigel (BD Biosciences) was added to each well of a 96-well plate and allowed to polymerize at 37°C for 1 h. Mutant or wild-type cells at first or second passage were harvested with trypsin/EDTA (BioWhittaker) and resuspended in DME/20% FBS. Cells (5,000/well) were then added to duplicate wells in 100 µl of growth medium in the presence or absence of recombinant FGF-2 (Invitrogen) or murine VEGF164 (Calbiochem). In a separate experiment, additional wells were seeded in the presence or absence of heparinase at 0.3 mU/ml (heparin lyase III; Seikagaku) and 10 ng/ml FGF-2. After 48 h, the degree of endothelial sprouting over the gel surface was measured by determining the net lengths of endothelial processes (in duplicate wells) viewed under phase-contrast light microscopy. The data is reported as fold stimulation relative to baseline growth without added growth factor. In some experiments, 100 µg/ml of unfractionated heparin (Scientific Protein Laboratories) was added to the wells during the sprouting period.
SDS-PAGE and Western blotting
Primary lung endothelial cells were seeded at equal density on gelatin-coated 12-well culture dishes immediately after harvest from the animals and allowed to grow to near confluence under standard culture conditions. After incubation for 4 h in serum-free DME without supplements, cells were exposed to FGF-2 or VEGF164 (10 ng/ml) for 10–60 min. Samples were solubilized and mixed with an equal volume of 125 mM Tris-HCl, pH 6.8, 4% SDS, 20% glycerol, 100 mM dithiothreitol, and 0.02% bromophenol blue. They were separated by electrophoresis in 4–15% gradient gels followed by electrotransfer to a nitrocellulose membrane, which was then probed using a polyclonal antibody (1:1,000; Cell Signaling Technology) that detects phosphorylated Thr202/Tyr204 forms of p44/42 MAPKs (Erk1/2). Bound antibody was detected using peroxidase-conjugated anti–rabbit antibody (1:2,000; Bio-Rad Laboratories) followed by visualization using SuperSignal chemiluminescent substrate for HRP detection (Pierce Chemical Co.). To ensure equal protein loading, membranes were stripped and reprobed using polyclonal antibodies against total Erk1/2. In a separate experiment, a subset of wells was treated with 0.5 mU/ml heparinase during the periods of serum starvation and growth factor stimulation. For Akt signaling, membranes were probed using 1:1,000 anti-phospho (Ser473)-Akt monoclonal antibody (Cell Signaling Technology). The bands were quantified by measurement of band intensity relative to background on a densitometer (GS800; Bio-Rad Laboratories).
Apoptosis assay on reconstituted extracellular matrix
Mutant cells were generated by transfection of subconfluent Ndst1f/fTekCre– primary murine lung endothelia with Adenoviral-Cre (Ad-Cre; UCSD Gene Therapy Core) at 109 pfu/ml for 1.5 h in normal culture growth medium. Infection with Adenoviral-GFP (Ad-GFP) was used as a control. Cells were added to Matrigel-coated wells in chamber slides. Permeabilization and TUNEL labeling were performed directly on chamber slides according to the manufacturer's instructions (In Situ Cell Death Detection kit; Roche), and samples were stained with Vector Blue Alkaline Phosphatase Substrate kit (Vector Laboratories) and nuclear fast red stain.
Statistics
Mean values of tumor volume, microvascular density on tumor sections or wound sections, microvascular skeletal length, caliber, grid intersection, and branching, and mean relative sprouting responses on extracellular matrix, were compared using one-tailed t tests. P < 0.05 was considered significant. Comparison of wound contraction rates in mutant versus wild-type mice was performed using a two-factor repeated-measures analysis of variance, testing for group differences in the change over time (SPSS Software v. 13.0).
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Acknowledgments |
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This work was supported by a U.S. Veteran's Administration Research Career Development Award and a Lopiccola Fellowship Award (M.M. Fuster), National Institutes of Health grants HD52920 and HL57345 (J.D. Esko), and the Infection and Wound Repair Core (supported through grants HL57345 [R.L. Gallo] and AI35796 [P. Sriramarao]).
The authors declare no competing financial interests.
Submitted: 18 October 2006
Accepted: 26 March 2007
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References |
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