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Am J Physiol Lung Cell Mol Physiol 288: L1117-L1123, 2005. First published February 18, 2005; doi:10.1152/ajplung.00370.2004
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Difference in proangiogenic potential of systemic and pulmonary endothelium: role of CXCR2

Aigul Moldobaeva and Elizabeth M. Wagner

Department of Medicine, Johns Hopkins University, Baltimore, Maryland

Submitted 4 October 2004 ; accepted in final form 11 February 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The systemic vasculature in and surrounding the lung is proangiogenic, whereas the pulmonary vasculature rarely participates in neovascularization. We studied the effects of the proangiogenic ELR+ CXC chemokine MIP-2 (macrophage inflammatory protein-2) on endothelial cell proliferation and chemotaxis. Mouse aortic, pulmonary arterial, and lung microvascular endothelial cells were isolated and subcultured. Proliferation ([3H]thymidine uptake) and migration (Transwell chemotaxis) were evaluated in each cell type at baseline and upon exposure to MIP-2 (1–100 ng/ml) without and with exposure to hypoxia (24 h)-reoxygenation. Baseline proliferation did not vary among cell types, and all cells showed increased proliferation after MIP-2. Aortic cell chemotaxis increased markedly upon exposure to MIP-2; however, neither pulmonary artery nor lung microvascular endothelial cells responded to this chemokine. Assessment of CXCR2, the G protein-coupled receptor through which MIP-2 signals, displayed no baseline difference in mRNA, protein, or cell surface expression among cell types. Exposure to hypoxia increased expression of CXCR2 of aortic endothelial cells only. Additionally, aortic cells, compared with pulmonary cells, showed significantly greater protein and activity of cathepsin S, a proteolytic enzyme important for cell motility. Thus the combined effects of increased cathepsin S activity, providing increased motility and enhanced CXCR2 expression after hypoxia, both contribute to the proangiogenic phenotype of systemic arterial endothelial cells.

angiogenesis; cathepsin S; hypoxia-reoxygenation; macrophage inflammatory protein-2

NEOVASCULARIZATION OF THE LUNG after chronic pulmonary embolism (5), interstitial pulmonary fibrosis (30), and bacterial infection (29) requires systemic vascular angiogenesis. New blood vessels invade the lung parenchyma from systemic bronchial arteries and/or intercostal arteries (5). Surprisingly, the pulmonary vasculature does not participate in the process and appears to be resistant to proliferation and neovascularization (19). This heterogeneity in endothelial cell responsiveness raises the question of why pulmonary arterial endothelium lacks proangiogenic potential in these pathological situations. A possible mechanism may be related to the prevailing ambient oxygen level. In systemic organs, vascular obstruction and ischemia result in systemic endothelial cell hypoxia. In general, tissue hypoxia increases the angiogenic potential of systemic arterial endothelial cells by inducing the upregulation of angiogenic factors and downregulation of antiangiogenic factors. However, an opposite situation exists for pulmonary arterial endothelium within the ventilated lung during pulmonary ischemia where normoxic or hyperoxic conditions prevail. In vitro assays of endothelial cell proliferation and migration have been performed exposing cells to a variety of proangiogenic factors, yet little attention has been given to examining the anatomically relevant endothelial cell subtype or appropriate ambient oxygen tension. There is increasing evidence for endothelial cell heterogeneity (7, 26), yet responsiveness to changes in ambient oxygen tensions likely has additional phenotypic affects on vital aspects of neovascularization such as endothelial cell proliferation and migration.

In previous studies, we showed that obstruction of the left pulmonary artery in mice induced the formation of a new vasculature that developed from intercostal arteries that crossed the pleural space and invaded the lung (20). These studies confirmed a process that has been identified in humans after chronic pulmonary thromboembolism as well as in models in several other species (3, 6, 11, 32) and provided an experimental model in which to study the progression of lung angiogenesis. Our subsequent studies established the importance of the Glu-Leu-Arg (ELR+) CXC chemokines in this angiogenesis model (25). These results provide further support for work by Arenberg and colleagues (2), who have shown that in several human disease states and animal models, the process of increased systemic vascularization of the lung involves these chemokines. The ELR+ CXC chemokines [IL-8; growth-related oncogene (GRO)-{alpha}, -{beta}, -{gamma}; ENA-78 (78-amino acid epithelial cell-derived neutrophil activator); GCP-2 (granulocyte chemotactic protein-2); NAP-2 (neutrophil-activating peptide-2)] have been implicated in the regulation of endothelial cell function including proliferation, migration, and tube formation (13, 14, 17, 27). The specific effects of ELR+ CXC chemokines on their target cells are mediated through binding CXCR1 and CXCR2 (1). Expression of CXCR2 on human endothelial cells has been confirmed in several studies (21) (22). In murine tissue, the ELR+ CXC ligands [macrophage inflammatory protein-2 (MIP-2), keratinocyte-derived chemokine, lipopolysaccharide inducible CXC chemokine] bind exclusively through CXCR2 (9). The effects of hypoxia on CXCR2 expression have not been determined. Thus we undertook the following series of experiments to test the hypothesis that the difference in proangiogenic potential of systemic endothelial cells relative to pulmonary endothelial cells is related to CXCR2 expression and influenced by ambient oxygen levels. We developed techniques to isolate and study primary cultures of mouse aortic, pulmonary artery, and lung microvascular endothelial cells. Specific experiments were designed to 1) confirm the presence of CXCR2 on mouse endothelial cells, 2) determine whether the difference in proangiogenic potential (proliferation and chemotaxis) of mouse systemic arterial endothelial cells and mouse pulmonary endothelial cells is determined by CXCR2 expression, 3) determine the effects of ambient oxygen conditions on CXCR2 expression, and 4) compare the expression of cathepsin S, a proteolytic enzyme important for endothelial cell migration and invasion (24), in arterial endothelial cells and pulmonary artery endothelial cells. Our results confirm that, relative to pulmonary artery endothelium, systemic arterial endothelial cells demonstrate increased responsiveness to a CXCR2 ligand, increased CXCR2 expression after hypoxia-reoxygenation, and increased cathepsin S activity. Furthermore, these results provide a mechanism for the lack of proangiogenic potential of pulmonary artery endothelium during inflammatory conditions that promote neovascularization.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Pulmonary artery and aortic endothelial cells. Procedures using animals were approved by the Johns Hopkins University Animal Care and Use Committee. The pulmonary artery and aorta were dissected from C57BL/6 mice (Jackson Laboratories, Bar Harbor, ME). The vessels were opened longitudinally and cut into two or three pieces. These pieces were placed with the intimal side down on Matrigel-coated, 35-mm tissue culture dishes in culture medium [2 ml of DMEM with 20% fetal calf serum (FCS), 150 µg/ml endothelial cell growth supplement, 100 µg/ml penicillin/streptomycin, 0.25 µg/ml amphotericin B, and 0.1 mM MEM with nonessential amino acids]. After 4–6 days, the pieces of vessels were removed and the endothelial cells that had migrated were treated with trypsin and replated to 0.2% gelatin-coated T-25 flasks. When confluent, endothelial cells were again transferred to T-75 flasks and subcultured. To confirm endothelial phenotypes, cells were immunostained for platelet endothelial cell adhesion molecule, von Willebrand's factor, and modified lipoprotein uptake (Dil-ac-LDL). Only cells with positive staining were used for subsequent experiments. All experiments were performed using endothelial cells isolated from seven mice and used between passages 2 and 10.

Lung microvascular endothelial cells. Mouse lung parenchyma was removed from a peripheral region devoid of large airways, rinsed with DMEM, minced, and digested in 1 ml of collagenase (1 mg/ml; Sigma, St. Louis, MO) at 37°C for 20 min with occasional agitation. The cellular digest was filtered through sterile mesh and centrifuged (400 g for 7 min). The cell pellet was resuspended (1 ml of complete medium) and plated on 0.2% gelatin-coated 24-well plates. After ~5–7 days, areas of cells exhibiting cobblestone morphology were selected, treated with trypsin, and replated to 0.2% gelatin-coated 24-well plates. When confluent, cells were transferred to T-25 flasks and subcultured.

Proliferation assay. Cellular DNA synthesis was assessed by [3H]thymidine uptake. Endothelial cells were maintained in medium with 5% FCS for 48 h, treated with trypsin, and seeded on 0.2% gelatin-coated 24-well plates (4 x 104 cells per well) in DMEM with 5% FCS and 5 µCI/ml [3H]thymidine. After 24 h, cells were washed and fixed (10 min on ice with 5% trichloroacetic acid), the DNA was released by alkaline lysis (0.5 N NaOH), and supernatants were quantified in a beta-counter (Beckmann LS6000SC; Beckman Coulter, Somerset, NJ).

Chemotaxis assay. Polycarbonate filters (5-µm pore size; Corning Costar, Cambridge, MA) were coated with 0.2% gelatin. Endothelial cells were detached with 2 mM EDTA/PBS, washed, and resuspended in DMEM with 0.1% BSA fatty acid free. 100 x 103 cells were added to the upper chamber and incubated (37°C, 1 h). After 1 h, medium in bottom chambers was replaced with DMEM with 0.1% BSA containing MIP-2 [1–100 ng/ml (27)] and incubated for 2 h. After 2 h, nonmigrated cells were removed with cotton tips, and migrated cells were fixed and stained using Diff-Quik stain set (Dade Behring, Newark, DE). Membranes were mounted on slides, and the average number of migrated cells was counted under a microscope (x20 objective)/six fields across each membrane.

Hypoxia exposure. Endothelial cells were loaded into a modular incubator chamber and flooded with a hypoxic gas mixture (4% O2, 5% CO2, and balance N2) for 24 h. Thereafter, cells were incubated in 95% air and 5% CO2 for 2 days before the experiment.

CXCR2 RNA isolation and RT-PCR. Total RNA was isolated from confluent endothelial cell cultures by a single-step guanidium thiocyanate/phenol-chloroform extraction using TRIzol reagent (Invitrogen, Carlsbad, CA). Using 100 ng of cDNA as a template, we performed quantification by an ABI Prism 5700 Sequence Detector (Applied Biosystems, Foster City, CA) using the TaqMan 5' nuclease activity from the TaqMan Universal PCR Master Mix, fluorogenic probes (Applied Biosystems), and oligonucleotide primers (Invitrogen). The mRNA expression levels of all samples were normalized to the levels of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) from the same sample. The primer sequence used for CXCR2 was: forward primer (5' GTG CCG CTG CTC ATC ATG 3'), reverse primer (5' AAG GAC GAC AGC GAA GAT GAC 3'), and TaqMan probe for GAPDH (5' TGC TAC GGG TTC ACA CTG CGC AC 3').

CXCR2 expression by flow cytometry. Endothelial cells were washed and detached with 2 mM EDTA/PBS and resuspended in PBS with 0.2% BSA. After being washed, cells were incubated with primary antibodies (anti-CXCR2 polyclonal antibodies, 1:10), washed, incubated with Alexa 488 anti-goat antibody (Molecular Probes, Eugene, OR), and analyzed. Mouse isotype IgG served as a negative control. Mean channel fluorescence for each experiment was normalized to normoxia controls.

CXCR2 and cathepsin S protein. Endothelial cells were treated with lysis buffer (1% SDS, 1 mM Na3Vo4, 5 mM NaF, and 5 µg/ml of leupeptin, aprotinin, and pepstatin) on ice. Lysates were boiled for 5 min, and total protein concentration was determined by bicinchoninic acid assay (Pierce, Rockford, IL). Protein (20 µg) was separated by SDS-PAGE, blotted on nitrocellulose, and blocked for 1 h at room temperature in 3% BSA/3% nonfat dry milk in Tris-buffered saline (50 mM Tris·HCl, pH 7.4) containing 0.1% Tween 20. After being washed in TBS-T, blots were incubated (1 h) with polyclonal rabbit anti-human antibodies to CXCR2 (Abcam, Cambridge, UK) or polyclonal goat anti-rat antibodies to cathepsin S (Santa Cruz Biotechnology, Santa Cruz, CA). Immunodetection was performed using horseradish peroxidase-conjugated anti-rabbit or anti-rat antibodies (Sigma) and enhanced chemiluminescence (Amersham Pharmacia Biotech, Arlington Heights, IL).

Cathepsin S activity. Specific catalytic activity of cathepsin S was determined fluorometrically by the hydrolysis of the synthetic substrate Z-VAL-VAL-Arg-MCA (Peptide Institute, Minoh, Osaka, Japan) according to Ref. 28 with the following modifications. After a 1-h incubation period with MIP-2 (10 ng/ml), cells were washed (3x) with cold PBS, suspended in extraction buffer (0.01% Triton X-100 in 0.1 M potassium phosphate buffer containing 1 mM EDTA, pH 7.5), and sonicated on ice. After centrifugation at 4°C (30 min at 14,000 rpm), the supernatant was removed for the cathepsin S activity assay. Supernatant (25 µl) was added to the buffer-activator solution (50 µl; 0.1 M potassium phosphate buffer, 5 mM EDTA, pH 7.5, and 5 mM DTT) and incubated (45 min, 40°C). After cooling, substrate (50 µl of 12.5 µM) was added to the mixture, and the increase in fluorescence measured (excitation 360 nm, emission 460 nm) after 30 min at 40°C. The calibration solution was prepared using 7-hydroxy-4-methylcoumarin (Molecular Probes). Cathepsin S activity was calculated as nmol of 7-hydroxy-4-methylcoumarin produced·mg protein–1·min–1. Cathepsin inhibitor II (10 µM; Z-FG-NHO-BzME; Calbiochem, San Diego, CA), an agent known to cause nonspecific cathepsin inhibition (4), was used to interfere with MIP-2-induced chemotaxis.

Statistics. Paired comparisons before and after treatment within a given endothelial cell type were evaluated with the Wilcoxon signed-rank test for paired comparisons. Unpaired comparisons were evaluated by the Mann-Whitney test. Comparisons across all cells types were made using the Kruskal-Wallis ANOVA, followed by Dunnett's multiple-comparison test. The n noted in RESULTS refers to the number of times the experiment was repeated on different cells, usually from different animals. Data are presented as means ± SE.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Endothelial cell proliferation. Basal proliferation did not differ among the three endothelial cell subtypes. However, each cell type showed a vigorous response to MIP-2 treatment. Although the specific concentration of MIP-2 (1–100 ng/ml) that resulted in the maximum increase in proliferation varied both within and among cell types, the average maximum proliferation was similar across cell types (252–258% of control, n = 5–6 experiments/cell type). Furthermore, the 24-h exposure to the hypoxic gas mixture did not alter basal proliferation evaluated 48 h after the exposure, nor did it alter the average maximum increase in proliferation observed after MIP-2 treatment in any cell type (213–277% of control).

Endothelial cell chemotaxis. Basal chemotaxis showed no difference across the three endothelial cell subtypes (n = 6 experiments/cell type, P = 0.0798). However, the response to MIP-2 demonstrated clear differences among cell types (Fig. 1, n = 7 experiments·cell type–1·concentration–1; P = 0.0004). Mouse aortic endothelial cells demonstrated a vigorous, dose-dependent, and significantly greater chemotaxis to MIP-2 than pulmonary endothelial cells (P < 0.01). Neither mouse pulmonary artery endothelial cells nor lung microvascular endothelial cells showed significant chemotaxis to the range of MIP-2 studied (1–100 ng/ml, P > 0.05). Each endothelial subtype demonstrated a trend toward increased chemotaxis after exposure to hypoxia alone (aortic endothelial cells: P = 0.059). The average effects of MIP-2 (1 ng/ml), hypoxia, and the combined effects of hypoxia and MIP-2 (1 ng/ml) are presented in Fig. 2, with individual experiments normalized to control responses observed during normoxia. Aortic cell responses to treatment were significantly greater than chemotaxis observed during basal conditions (P = 0.0098). However, neither pulmonary artery endothelial cells (P = 0.661) nor lung microvascular cells (P = 0.8990) displayed differences from basal chemotaxis. In aortic endothelial cells, average responses to hypoxia and MIP-2 (1 ng/ml) individually appeared approximately additive to the experimental condition of MIP-2 after hypoxia. However, due to inherent variability this result did not reach statistical significance (P = 0.6277). When comparing chemotaxis in aortic cells exposed to both hypoxia and MIP-2 with the sum of the individual effect of hypoxia plus the individual effect of MIP-2, we found the changes to be similar (P = 0.85). This analysis of individual experiments suggests that the responses to MIP-2 and hypoxia were additive.



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  		Fig. 1. Percent change in chemotaxis from baseline levels in response to macrophage inflammatory protein (MIP-2, 1–100 ng/ml) in aortic (ao), pulmonary artery (pa), and lung microvascular (lmv) endothelial cells. Only ao endothelial cells showed a significant increase in chemotaxis in response to MIP-2 (n = 7 experiments·cell type–1·concentration–1, P = 0. 0004).

 


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  		Fig. 2. Comparison of endothelial responsiveness to MIP-2 (1 ng/ml), hypoxia, and hypoxia before MIP-2 (1 ng/ml) treatment in ao, pa, and lmv endothelial cells. Only ao cells showed a significant increase in chemotaxis with treatments (P = 0.0098).

 
CXCR2 expression. To confirm the presence of CXCR2 on the three endothelial cell subtypes, we measured mRNA and CXCR2 protein and cell surface expression by fluorescence-activated cell sorting analysis. As shown in Fig. 3, basal expression of CXCR2 mRNA normalized to GAPDH was equivalent among cell types (P = 0.9062, n = 5–7 experiments/cell type). After exposure to hypoxia followed by reoxygenation, there was a significant increase in the CXCR2 expression in aortic endothelial cells only (P = 0.0180). A representative Western blot of total CXCR2 protein in endothelial cell lysates using polyclonal anti-CXCR2 antibody demonstrated a specific band (55 kDa) corresponding with CXCR2 in cell lysates from all three populations of endothelial cells (Fig. 4). Basal protein levels of CXCR2 were equivalent among cell types (P = 0.6703, n = 3 experiments/group). Only aortic endothelial cells consistently in each replicate experiment showed an increase in CXCR2 protein after hypoxia-reoxygenation. Results from the assessment of cell surface receptors using polyclonal anti-CXCR2 are shown in Fig. 5 (n = 6–7 experiments/group). Mean fluorescence from anti-CXCR2 binding did not vary among cell types during basal, normoxic conditions (P = 0.9062). However, the expression of CXCR2 receptors of aortic endothelial cells only was significantly enhanced after hypoxia-reoxygenation (P = 0.0180).



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  		Fig. 3. CXCR2 mRNA levels in ao, pa, and lmv endothelial cells. There were no differences in baseline level of mRNA normalized to GAPDH in the endothelial cell subtypes. Exposure to hypoxia caused a significant increase in mRNA CXCR2 in ao cells only (*P = 0.0180, n = 5–7 experiments/group).

 


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  		Fig. 4. Representative Western blot of CXCR2 protein in ao, pa, and lmv endothelial cells.

 


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  		Fig. 5. CXCR2 cell surface expression of the 3 endothelial cell subtypes. No difference in baseline expression was observed (P = 0.9062). Only ao endothelial cells showed a significant increase in CXCR2 surface expression after hypoxia (*P = 0.0180, n = 6–7 experiments/group).

 
Cathepsin S activity. Based on pilot microarray data (not shown) and the work of others (24), we studied protein expression and activity of the lysomal, proteolytic enzyme cathepsin S in aortic and pulmonary artery endothelial cells. These two endothelial cell subtypes were selected because they were expected to represent homogeneous populations, whereas lung microvascular cells likely represent a more heterogeneous population. Figure 6A shows a representative Western blot displaying distinct bands representative of cathepsin S at 28 kDa. Quantification of pixel intensity normalized to {beta}-actin demonstrated a small but significant difference between the level of cathepsin S protein of aortic endothelial cells compared with pulmonary artery endothelial cells after MIP-2 exposure (Fig. 6B; P = 0.0079, n = 5 experiments/group). As shown in Fig. 7, overall cathepsin S activity in aortic cells was significantly greater than in pulmonary artery endothelial cells (P = 0.0036). Specifically, MIP-2 significantly increased cathepsin S activity in aortic cells relative to pulmonary artery endothelial cells (P = 0.0087, n = 6 experiments/cell type). The combined effects of hypoxia-reoxygenation and MIP-2 further enhanced cathepsin S activity in aortic cells (P < 0.05, n = 3 experiments/cell type). Additional experiments in aortic endothelial cells (Fig. 8, n = 4 experiments) demonstrated the effects of cathepsin II inhibitor on MIP-2-induced chemotaxis. Chemotaxis was completely blocked by this cathepsin S inhibitor (P = 0.02).



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  		Fig. 6. A: representative Western blot of cathepsin S protein expression in ao and pa endothelial cells. B: incubation with MIP-2 (10 ng/ml) caused a significant difference in cathepsin S protein between ao and pa endothelial cells (P = 0.0079, n = 5 experiments/group).

 


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  		Fig. 7. Comparison of the effects of MIP-2 (10 ng/ml), hypoxia, and hypoxia + MIP-2 on cathepsin S activity in ao and pa endothelial cells. Overall cathepsin S activity in ao cells was significantly greater than in pa endothelial cells (P = 0.0036). MIP-2 elicited a significant increase in cathepsin S activity in ao cells relative to pulmonary cells (P = 0.0087, n = 6 experiments/cell type). The combined effects of hypoxia-reoxygenation and MIP-2 further enhanced cathepsin S activity in ao cells (P < 0.05, n = 3 experiments).

 


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  		Fig. 8. Average effects of cathepsin II inhibitor on MIP-2-induced chemotaxis of ao endothelial cells (n = 4 experiments). Chemotaxis was completely blocked by this cathepsin S inhibitor (*P = 0.02).

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Increased attention has focused recently on the heterogeneity of endothelial cells and the unique attributes of endothelial cells from different organs as well as from different locations within a given vasculature (7, 26). Emerging observations demonstrate that endothelial cells have specialized properties, capabilities, and molecular signaling that appear to be dependent on their anatomical origin. Perhaps one of the most perplexing differences in endothelial cell potential is the long-standing observation that the systemic vasculature within and surrounding the lung responds vigorously to pulmonary vascular ischemia by sending new vessels to the lung parenchyma while the pulmonary endothelium remains unresponsive. In a broad spectrum of diseases including asthma, cystic fibrosis, interstitial pulmonary fibrosis, lung carcinoma, and chronic thromboembolic disease, the unique capacity of systemic vessels to proliferate and to invade ischemic lung tissues has been observed. In this study, we sought to determine factors that distinguish systemic arterial endothelium from pulmonary artery endothelium in defining a proangiogenic phenotype. Additionally, the importance of CXC chemokines in promoting systemic neovascularization in the lung has been reported for several different angiogenesis models (2, 12, 25). Therefore, in the reported series of experiments, we focused on determining the differences in systemic arterial and pulmonary endothelial cells with regard to CXC chemokine responsiveness and CXC receptor expression. In mice, MIP-2 is one of the most potent of the proangiogenic ELR+ CXC chemokine ligands, which binds to the G protein-coupled receptor CXCR2. Although mice have been shown to lack CXCR1, in other species, CXCR1 and CXCR2 have been shown to account for all ELR+ CXC chemokine signaling (9). Because chemokines typically elicit activation and migration of their target cells in vivo, we compared proliferation and chemotaxis of three endothelial cell subtypes in vitro. We found that endothelial cell chemotaxis was markedly enhanced in systemic arterial endothelial cells compared with either pulmonary artery or lung microvascular endothelial cells in response to MIP-2. This response was observed at all doses of MIP-2 studied (Fig. 1). Strieter and colleagues (27) showed a somewhat greater chemotactic response of human adrenal gland capillary endothelial cells to the human analog of MIP-2, GRO-{beta},{gamma} averaging 3.5- to 4.5-fold increase at a chemokine concentration of 10 ng/ml. These observations further support the overall heterogeneity of endothelial cell responsiveness. Interestingly, however, in the present study, equivalent levels of CXCR2 expression among the three endothelial cell subtypes was confirmed by measurement of CXCR2 mRNA, protein, and cell surface receptor activity (Figs. 3–5). Thus enhanced chemotaxis of systemic arterial endothelial cells in the presence of MIP-2 could not be attributed to increased receptor expression. Furthermore, although systemic arterial endothelial cell chemotaxis differed markedly in response to MIP-2, basal chemotaxis did not differ, and MIP-2 was equally effective at increasing endothelial cell proliferation among all three endothelial cell subtypes. Overall these results suggest that the proangiogenic influence of MIP-2 on aortic endothelial cells is related to cell motility.

Endothelial cell migration is central to the overall process of angiogenesis and requires the proteolytic modification of extracellular matrix. Many studies have focused on the importance of the matrix metalloproteases and serine proteases in permitting endothelial cells to bore into surrounding interstitium and form new vessels (31). Several recent studies have focused on the lysosomal cysteine proteases, or cathepsins, in supporting a similar role in tumorigenesis (8, 10, 15). A recent study specifically demonstrated a role for cathepsin S in mediating angiogenesis (24). Cathepsin S is capable of degrading extracellular matrix proteins such as laminin, collagens, elastin, and chondroitin sulphate proteoglycan. Shi and colleagues (24) showed that inflammatory mediators (TNF-{alpha}, IFN-{gamma}, IL-1{beta}) and growth factors (VEGF, bFGF) upregulate cathepsin S expression in endothelial cells. Additionally, endothelial cells recovered from cathepsin S –/– mice showed impaired invasion in Matrigel and collagen gel assays. These properties of inducibility and the capacity to degrade extracellular matrix are consistent with the requirements of endothelial cells during neovascularization. An initial preliminary screening experiment using microarray analysis suggested cathepsin expression might differ among endothelial cell subtypes. We focused our experiments on primary cultures of the two most homogeneous endothelial cell populations, those of systemic aortic endothelial cells and pulmonary arterial endothelial cells, to determine whether the marked differences in chemotaxis to MIP-2 might be related to genotypic differences in cathepsin S. MIP-2 caused a significant increase in cathepsin S protein expression in aortic endothelial cells relative to pulmonary artery endothelium. More impressive, however, was the increase in cathepsin S activity in aortic relative to pulmonary artery endothelial cells after MIP-2 treatment (Fig. 7). These results are consistent with the hypothesis that cathepsin S activity modulates chemotaxis in aortic but not pulmonary artery endothelial cells. By use of a nonspecific cathepsin inhibitor, chemotaxis induced by MIP-2 was largely prevented. This result lends additional support for the overall importance of cathepsins for systemic endothelial cell migration.

We also studied the effects of hypoxia-reoxygenation on proliferation and chemotaxis in the three endothelial cell subtypes. Tissue hypoxia is an important stimulus and regulator for new vessel growth in most systemic organs. Hypoxia can increase the angiogenic potential of endothelial cells by producing an oxidative stress. Brief exposure to hypoxia (30–60 min) followed by reoxygenation significantly accelerated (threefold) the rate of tubular morphogenesis (16). Additionally, Maulik and Das (18) showed that human coronary endothelial cells plated on Matrigel after hypoxia-reoxygenation showed increased tubular morphogenesis. Thus we questioned whether a change in the ambient oxygen conditions might alter the responsiveness of the endothelial cells studied. Specifically, since pulmonary artery endothelium is normally exposed to mixed venous PO2, we questioned whether the in vitro culture conditions routinely used predispose these cells to artificial responses. Our choice of a 24-h exposure to hypoxia followed by 48 h of reoxygenation was based on experimental constraints and preliminary studies. Different results might be obtained if cells were exposed to a shorter/longer time course of hypoxia-reoxygenation or lesser/greater level of hypoxia. However, we selected this PO2 based on the normal environment of pulmonary artery endothelial cells hoping to mimic in vivo oxygen conditions. We did not see significantly different responses in pulmonary endothelial cells after hypoxia-reoxygenation compared with the normoxia controls. The 24-h exposure may have been insufficient to cause lasting changes in these cells. A persistent hypoxic growth environment may be required to fully restore in vivo attributes to pulmonary artery endothelial cells. Conversely, the effects of hypoxia on systemic arterial endothelial cells more accurately mimic in vivo conditions where systemic tissue hypoxia followed by reoxygenation is representative of in vivo systemic ischemia. Our results showed that CXCR2 expression was sensitive to hypoxia-reoxygenation only in aortic endothelial cells (Figs. 3–5). CXCR2 mRNA, protein, and cell surface expression were significantly increased after hypoxia-reoxygenation. These results could account for an enhanced responsiveness of systemic arterial endothelial cells to ELR+ CXC chemokines after an exposure to hypoxia. Furthermore, our results demonstrated that the effects of MIP-2 and hypoxia-reoxygenation appeared to be additive. Previous work by Schioppa and colleagues (23) showed the upregulation of another chemokine receptor CXCR4 in human umbilical vein endothelial cells after exposure to hypoxia (1% O2, 24 h) and increased chemotaxis to the chemokine ligand stromal-derived factor 1 (CXCL12). Overall, these results demonstrate selective induction of chemokine receptors exposed to low oxygen tensions and the importance of ambient oxygen tensions in assessing angiogenic properties of endothelial cells.

In summary, we have shown that systemic arterial endothelial cells are more responsive to the ELR+ C-X-C chemokine MIP-2 than pulmonary endothelial cells. Under normoxic, control conditions, the increased migration observed in arterial endothelial cells could not be accounted for by differences in chemokine receptor levels. However, the increased responsiveness is due, in part, by enhanced cathepsin S activity in arterial endothelial cells. Additionally, hypoxia-reoxygenation causes an increase in CXCR2 expression in aortic endothelial cells that contributes to an enhanced proangiogenic phenotype.


   FOOTNOTES
 

Address for reprint requests and other correspondence: E. M. Wagner, Johns Hopkins Asthma and Allergy Center, Div. of Pulmonary and Critical Care Medicine, 5501 Hopkins Bayview Circle, Baltimore, MD 21224 (E-mail: wagnerem{at}jhmi.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.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Addison CL, Daniel TO, Burdick MD, Liu H, Ehlert JE, Xue YY, Buechi L, Walz A, Richmond A, and Strieter RM. The CXC chemokine receptor 2, CXCR2, is the putative receptor for ELR+ CXC chemokine-induced angiogenic activity. J Immunol 165: 5269–5277, 2000.[Abstract/Free Full Text]
  2. Arenberg DA, Keane MP, DiGiovine B, Kunkel SL, Morris SB, Xue YY, Burdick MD, Glass MC, Iannettoni MD, and Strieter RM. Epithelial-neutrophil activating peptide (ENA-78) is an important angiogenic factor in non-small cell lung cancer. J Clin Invest 102: 465–472, 1998.[ISI][Medline]
  3. Charan NB and Carvalho P. Angiogenesis in bronchial circulatory system after unilateral pulmonary artery obstruction. J Appl Physiol 82: 284–291, 1997.[Abstract/Free Full Text]
  4. Demuth HU, Schierhorn A, Bryan P, Hofke R, Kirschke H, and Bromme D. N-peptidyl, O-acyl hydroxamates: comparison of the selective inhibition of serine and cysteine proteinases. Biochim Biophys Acta 1295: 179–186, 1996.[CrossRef][Medline]
  5. Endrys J, Hayat N, and Cherian G. Comparison of bronchopulmonary collaterals and collateral blood flow in patients with chronic thromboembolic and primary pulmonary hypertension. Heart 78: 171–176, 1997.[Abstract/Free Full Text]
  6. Fadel E, Mazmanian GM, Chapelier A, Baudet B, Detruit H, de Montpreville V, Libert JM, Wartski M, Herve P, and Dartevelle P. Lung reperfusion injury after chronic or acute unilateral pulmonary artery occlusion. Am J Respir Crit Care Med 157: 1294–1300, 1998.[Abstract/Free Full Text]
  7. Gebb S and Stevens T. On lung endothelial cell heterogeneity. Microvasc Res 68: 1–12, 2004.[CrossRef][ISI][Medline]
  8. Gondi CS, Lakka SS, Yanamandra N, Olivero WC, Dinh DH, Gujrati M, Tung CH, Weissleder R, and Rao JS. Adenovirus-mediated expression of antisense urokinase plasminogen activator receptor and antisense cathepsin B inhibits tumor growth, invasion, and angiogenesis in gliomas. Cancer Res 64: 4069–4077, 2004.[Abstract/Free Full Text]
  9. Horuk R. Chemokine receptors. Cytokine Growth Factor Rev 12: 313–335, 2001.[CrossRef][ISI][Medline]
  10. Joyce JA, Baruch A, Chehade K, Meyer-Morse N, Giraudo E, Tsai FY, Greenbaum DC, Hager JH, Bogyo M, and Hanahan D. Cathepsin cysteine proteases are effectors of invasive growth and angiogenesis during multistage tumorigenesis. Cancer Cell 5: 443–453, 2004.[CrossRef][ISI][Medline]
  11. Karsner H and Ghoreyeb A. Studies in infarction: the circulation in experimental pulmonary embolism. J Exp Med 18: 507–522, 1913.[Abstract/Free Full Text]
  12. Keane MP, Belperio JA, Moore TA, Moore BB, Arenberg DA, Smith RE, Burdick MD, Kunkel SL, and Strieter RM. Neutralization of the CXC chemokine, macrophage inflammatory protein-2, attenuates bleomycin-induced pulmonary fibrosis. J Immunol 162: 5511–5518, 1999.[Abstract/Free Full Text]
  13. Keane MP and Strieter RM. The role of CXC chemokines in the regulation of angiogenesis. Chem Immunol 72: 86–101, 1999.[Medline]
  14. Koch AE, Polverini PJ, Kunkel SL, Harlow LA, DiPietro LA, Elner VM, Elner SG, and Strieter RM. Interleukin-8 as a macrophage-derived mediator of angiogenesis. Science 258: 1798–1801, 1992.[Abstract/Free Full Text]
  15. Lakka SS, Gondi CS, Yanamandra N, Olivero WC, Dinh DH, Gujrati M, and Rao JS. Inhibition of cathepsin B and MMP-9 gene expression in glioblastoma cell line via RNA interference reduces tumor cell invasion, tumor growth and angiogenesis. Oncogene 23: 4681–4689, 2004.[CrossRef][ISI][Medline]
  16. Lelkes PI, Hahn KL, Sukovich DA, Karmiol S, and Schmidt DH. On the possible role of reactive oxygen species in angiogenesis. Adv Exp Med Biol 454: 295–310, 1998.[ISI][Medline]
  17. Li A, Dubey S, Varney ML, Dave BJ, and Singh RK. IL-8 directly enhanced endothelial cell survival, proliferation, and matrix metalloproteinases production and regulated angiogenesis. J Immunol 170: 3369–3376, 2003.[Abstract/Free Full Text]
  18. Maulik N and Das DK. Redox signaling in vascular angiogenesis. Free Radic Biol Med 33: 1047–1060, 2002.[CrossRef][ISI][Medline]
  19. Milne EN and Zerhouni EA. Blood supply of pulmonary metastases. J Thorac Imaging 2: 15–23, 1987.[Medline]
  20. Mitzner W, Lee W, Georgakopoulos D, and Wagner E. Angiogenesis in the mouse lung. Am J Pathol 157: 93–101, 2000.[Abstract/Free Full Text]
  21. Murdoch C, Monk PN, and Finn A. Cxc chemokine receptor expression on human endothelial cells. Cytokine 11: 704–712, 1999.[CrossRef][ISI][Medline]
  22. Salcedo R, Resau JH, Halverson D, Hudson EA, Dambach M, Powell D, Wasserman K, and Oppenheim JJ. Differential expression and responsiveness of chemokine receptors (CXCR1–3) by human microvascular endothelial cells and umbilical vein endothelial cells. FASEB J 14: 2055–2064, 2000.[Abstract/Free Full Text]
  23. Schioppa T, Uranchimeg B, Saccani A, Biswas SK, Doni A, Rapisarda A, Bernasconi S, Saccani S, Nebuloni M, Vago L, Mantovani A, Melillo G, and Sica A. Regulation of the chemokine receptor CXCR4 by hypoxia. J Exp Med 198: 1391–1402, 2003.[Abstract/Free Full Text]
  24. Shi GP, Sukhova GK, Kuzuya M, Ye Q, Du J, Zhang Y, Pan JH, Lu ML, Cheng XW, Iguchi A, Perrey S, Lee AM, Chapman HA, and Libby P. Deficiency of the cysteine protease cathepsin S impairs microvessel growth. Circ Res 92: 493–500, 2003.[Abstract/Free Full Text]
  25. Srisuma S, Biswal SS, Mitzner WA, Gallagher SJ, Mai KH, and Wagner EM. Identification of genes promoting angiogenesis in mouse lung by transcriptional profiling. Am J Respir Cell Mol Biol 29: 172–179, 2003.[Abstract/Free Full Text]
  26. Stevens T, Rosenberg R, Aird W, Quertermous T, Johnson FL, Garcia JG, Hebbel RP, Tuder RM, and Garfinkel S. NHLBI workshop report: endothelial cell phenotypes in heart, lung, and blood diseases. Am J Physiol Cell Physiol 281: C1422–C1433, 2001.[Abstract/Free Full Text]
  27. Strieter RM, Polverini PJ, Kunkel SL, Arenberg DA, Burdick MD, Kasper J, Dzuiba J, Van Damme J, Walz A, Marriott D, Chan SY, Roczniak S, and Shanafelt AB. The functional role of the ELR motif in CXC chemokine-mediated angiogenesis. J Biol Chem 270: 27348–27357, 1995.[Abstract/Free Full Text]
  28. Sugano E, Tomita H, Abe T, Yamashita A, and Tamai M. Comparative study of cathepsins D and S in rat IPE and RPE cells. Exp Eye Res 77: 203–209, 2003.[CrossRef][ISI][Medline]
  29. Thurston G, Murphy TJ, Baluk P, Lindsey JR, and McDonald DM. Angiogenesis in mice with chronic airway inflammation: strain-dependent differences. Am J Pathol 153: 1099–1112, 1998.[Abstract/Free Full Text]
  30. Turner-Warwick M. Precapillary systemic-pulmonary anastomoses. Thorax 18: 225–237, 1963.[Medline]
  31. Visse R and Nagase H. Matrix metalloproteinases and tissue inhibitors of metalloproteinases: structure, function, and biochemistry. Circ Res 92: 827–839, 2003.[Abstract/Free Full Text]
  32. Weibel ER. Early stages in the development of collateral circulation to the lung in the rat. Circ Res 8: 353–376, 1960.[Abstract/Free Full Text]



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