Published online before print August 19, 2004, doi: 10.1161/01.ATV.0000142810.27849.8f
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
© 2004 American Heart Association, Inc.
Vascular Biology |
Human Endothelial Progenitor Cells Tolerate Oxidative Stress Due to Intrinsically High Expression of Manganese Superoxide Dismutase
From the Departments of Anesthesiology (T.H., T.E.P., Z.S.K.), Molecular Pharmacology and Experimental Therapeutics (A.T., Z.S.K.), and Internal Medicine (E.L.H., A.T., N.M.C.), Division of Cardiovascular Disease, Mayo Clinic College of Medicine, Rochester, Minn; and the Free Radical and Radiation Biology Program (L.W.O.), Department of Radiation Oncology, The University of Iowa, Iowa City, Iowa.
Correspondence to Zvonimir S. Katusic, Mayo Clinic, Joseph Building 4-184, 200 First Street SW, Rochester, MN 55905. E-mail katusic.zvonimir{at}mayo.edu
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
Objective— Endothelial progenitor cells (EPCs) display a unique aptitude to promote angiogenesis and restore endothelial function of injured vessels. How progenitor cells can execute a regenerative program in the unfavorable environment of injury/inflammation-induced oxidative stress is poorly understood. We hypothesized that EPCs are resistant to oxidative stress and that this resistance is due to high expression and activity of antioxidant enzymes.
Methods and Results— EPCs outgrown from human blood of healthy subjects demonstrated a marked resistance to cytotoxic effect of LY83583 (an 
generator), tumor necrosis factor-
, and serum depletion. LY83583 inhibited in vitro tube formation by human umbilical vein endothelial cells (HUVECs) and human coronary artery endothelial cells (CAECs), but not by EPCs. Compared with HUVECs and CAECs, EPCs exhibited 
3- to 4-fold higher expression and activity of manganese superoxide dismutase (MnSOD), but not copper zinc superoxide dismutase (CuZnSOD) or catalase. The antioxidant profile in EPCs was associated with preservation of the mitochondrial network when exposed to LY83583. Moreover, cytotoxic effects of LY83583 on CAECs and HUVECs were reversed by adenoviral overexpression of MnSOD.
Conclusions— Human EPCs are resistant to oxidative stress. High intrinsic expression of MnSOD is a critical mechanism protecting EPCs against oxidative stress.
We provide evidence that human endothelial progenitor cells (EPCs) are resistant to oxidative stress imposed by induced concentration of superoxide anions. Analysis of this phenomenon demonstrated that high intrinsic expression of manganese superoxide dismutase in EPCs is a critical mechanism underlying EPCs resistance to oxidative stress.
Key Words: endothelium • antioxidants • angiogenesis • superoxide • nitric oxide synthase • endothelial progenitor cells
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Evidence continues to accumulate on the importance of endothelial progenitor cells (EPCs) in neovascularization of ischemic tissues and inhibition of neointimal proliferation after balloon-injury.1–5 EPCs isolated from circulating blood may originate from bone marrow6,7 or from resident EPCs embedded within organs.8 In contrast to differentiated mature endothelial cells, EPCs have a high proliferation potential and can be expanded extensively in vitro.6,7,9 Transplantation of EPCs enhances vascular development by in situ differentiation and proliferation within ischemic organs.10,11 Successful transplantation and beneficial therapeutic effect of EPCs in ischemic tissues of experimental animal models1,2 and in humans4,12 suggest that EPCs may exhibit a high survival potential under unfavorable conditions of ischemia-reperfusion and associated oxidative stress.
See page 1977
Excessive production of reactive oxygen species (ROS) and reactive nitrogen oxide species is an essential mechanism underlying pathogenesis of endothelial dysfunction and vascular disease.13 Although it has been established that antioxidant enzymes, namely superoxide dismutases (SODs, converting to H2O2), catalase, and peroxidases (H2O2 scavengers), are critical in the defense against oxidative stress, 14 the antioxidant profile of progenitor cells remains poorly understood. Here we demonstrate that human EPCs possess a unique property to withstand oxidative injury and that elevated expression of manganese superoxide dismutase (MnSOD, a mitochondria-located SOD) is a critical intrinsic mechanism protecting EPCs against oxidative stress.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
For expanded Methods, please see the online section, available at http://atvb.ahajournals.org.
EPC Isolation, Cell Culture, and Phenotyping
The protocol for collection and use of human blood samples was approved by the Institutional Review Board at the Mayo Clinic. EPCs from blood of 15 healthy volunteers were obtained by density gradient centrifugation with Ficoll-Paque Plus (Amersham Biosciences Corp) as previously described.7,9 Isolated mononuclear cells were plated at a density of 2x107 cells per well on 6-well plates coated with human fibronectin (R&D Systems Inc) in endothelial growth medium-2 (EGM-2, Cambrex Corp), composed of endothelial cell basal medium-2 (EBM-2), 5% fetal bovine serum, and growth factors. Cell colonies appeared at 2 to 3 weeks. At 4 weeks, subconfluent cell colonies were passaged and cells were subsequently cultured in EGM-2. In parallel, human umbilical vein endothelial cells (HUVECs; Clonetics) and human coronary artery endothelial cells (CAECs; Clonetics) were cultured in EGM-2 with the same list of additives as used for EPCs. The majority of the experiments were performed on the cells cultured from passages 4 to 8. In some control experiments, studies were performed on the cells of passages 2 to 9. All cells studied were at similar confluence within the same experiment. Morphological appearance, indirect immunofluorescence, and flow cytometry analysis were used to define endothelial cell phenotype of EPCs as previously described.6,7,9,15 For detection of von Willebrand factor and Flk1, cells were permeabilized by incubation with 0.1% Triton X-100 for 2 minutes.7
Western Blot Analysis
Western blotting was performed as previously described.16 Rabbit polyclonal antibodies against MnSOD,17 CuZnSOD,17 cyclooxygenase-2 (COX-2; Cayman Chemical), and vascular cell adhesion molecule-1 (VCAM-1; Santa Cruz Biotechnology Inc) were used at dilutions of 1:1000, 1:500, 1:250, and 1:500, respectively; mouse monoclonal antibodies against catalase (Sigma-Aldrich Co) at 1:300 and endothelial nitric oxide synthase (eNOS, BD Biosciences) at 1:300. Blots probed with actin (1:500, Santa Cruz Biotechnology Inc) were used as loading controls. In some experiments, cells were treated with a cytokine cocktail (30 U/mL interlukin-1 [Sigma-Aldrich Co]+1 µg/mL tumor necrosis factor- [TNF-; R&D Systems Inc]+300 U/mL interferon-
[Sigma-Aldrich Co]) or TNF- alone for 24, 48, and 72 hours. The optical density of the band was measured by using Scion Image (Scion Corp). Protein expression was normalized to actin and expressed as relative densitometric units.
Assessment of Cell Viability
EPCs, CAECs, and HUVECs, plated at 2x104 per well on 24-well plates (3 wells for each sample), were treated with 1, 4, or 8 µmol/L LY83583 (Biomol Research Labs Inc) or 0.1 ng/mL TNF- for 72 hours, and medium along with detached cells removed. Remaining attached cells were harvested by trypsinization and counted in a hemocytometer. Experiments were performed 3 to 6 times. The treated cell numbers were normalized to % of untreated control (cells cultured in EGM-2 alone for 72 hours).
In separate experiments, HUVECs were incubated with replication-deficient adenoviral constructs containing human MnSOD (AdMnSOD, ViraQuest Inc) at 50 multiplicities of infection (MOI) in 1 mL of EGM-2 for 24 hours. Cells were then recovered in fresh EGM-2 for another 24 hours. For controls, cells were transduced with 50 MOI adenovirus expressing ß-galactosidase (AdLacZ) for 24 hours. HUVECs were then treated with 10 µmol/L LY83583 for 72 hours. The attached cells were harvested by trypsinization and assayed for cell viability by trypan blue (Sigma-Aldrich Co) dye exclusion.
Terminal Deoxynucleotidyl Transferase-Mediated dUTP Nick-End Labeling (TUNEL) Assay
Cells (5x104 per well) were seeded on 2-well chamber slides. After treatment with LY83583 (for 72 hours), cisplatin (for 24 hours), or EBM-2 (for 24 hours), DNA strand breaks were labeled with fluorescien-12-dUTP. TUNEL-positive cells (green-staining nuclei) were counted from 15 random fields of each sample using a fluorescent microscopy; data were expressed as percentage of total number of cells.
In Vitro Tube Formation Assay
Endothelial tube formation was assessed using Matrigel assay (BD Biosciences). Cells were seeded with a density of 7x104/well on 24-well plates (3 wells for each condition) coated with 250 µL Matrigel and incubated with EGM-2 alone or EGM-2 supplemented with 8 µmol/L LY83583 for 10 hours at 37°C. Tube formation was examined by a phase-contrast microscopy and tube circles were counted in the center field (x4 magnification) of each well. Four independent experiments were performed.
Tracking Mitochondrial Integrity
Fluorescent confocal microscopy and electron microscopy were used to monitor morphology of mitochondria. HUVECs, CAECs, and EPCs grown on 2-well chamber slides were treated with generating compound LY83583 (8 µmol/L) for 24 hours (in some CAECs groups, cells were transduced with 50 MOI AdMnSOD or AdLacZ for 24 hours and recovered in fresh EGM-2 for another 24 hours before LY83583 treatment). Cells were then loaded with MitoTracker Red CM-H2XRos (100 nM for 1 hour, Molecular Probes Inc), a marker of mitochondrial integrity. Cells were then fixed and costained with the nuclear fluorescent dye Hoescht 33528 and examined using fluorescent confocal microscopy. For quantification of the cells with bright punctate-like mitochondrial appearance, 223±14 cells from 10 to 15 random fields were counted for each sample. In addition, electron microscopy (1200 EXII, JEOL) was used, as described,18 to capture changes in mitochondrial structure at high-resolution after oxidative stress with LY83583 (10 µmol/L for 24 hours).
Zymograms of Antioxidant Enzymes
Total protein of each sample (100 µg per well) was separated in a native polyacrylamide gel. The enzymatic activity staining experiments for SOD and catalase were performed as described by Beauchamp and Fridovich19 and Sun et al,20 respectively. The optical densities of the bands were measured by Scion Image (Scion Corp).
Statistical Analysis
Data are presented as mean±SD. Differences between mean values of multiple groups were analyzed using ANOVA followed by Bonferroni t test multiple comparison procedure (SigmaStat 2.03 for Windows). Comparison between 2 groups was made using Student t test. P<0.05 was considered statistically significant.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Characterization of EPCs Phenotype
Two to 15 colonies per blood sample of healthy individuals were grown 2 to 3 weeks after isolation of mononuclear cells (Figure IA, available online at http://atvb.ahajournals.org). At confluence, EPCs exhibited the "cobblestone" morphology and monolayer growth pattern typical of endothelial cells (Figure IB). The endothelial phenotype of the outgrown EPCs were further characterized by positive staining for acetylated low-density lipoprotein uptake, isolectin binding, and expression of endothelial markers such as Flk-1, von Willebrand factor , CD31, and VE-Cadherin (Figure IC through IE, IG, and IH). In addition, EPCs demonstrated expression of the endothelial-specific eNOS, though at reduced levels compared with mature endothelial cells, HUVECs, and CAECs (Figure 1). Typical for endothelial cells, challenge with a cytokine cocktail for 24 and 48 hours decreased eNOS protein expression in both HUVECs and EPCs, yet with a lower eNOS level than found in EPCs.
|
P<0.05, compared with EPCs with cytokines for 24 or 48 hours.
Comparison of Survival Ability and Function of EPCs and Mature Endothelial Cells Under Oxidative Stress
Human EPCs displayed high tolerance to cytotoxicity induced by napthoquinolinedione LY83583, an established generator of intracellular ,21 and by conditions shown to induce oxidative stress (TNF- and serum depletion).22,23 Incubation with 1, 4, or 8 µmol/L LY83583 or 0.1 ng/mL TNF- for 72 hours significantly decreased the number of attached HUVECs and CAECs, as compared with EPCs (Figure 2). In fact, as shown in Figure 2A, severe cytotoxicity, which included cell deformation, detachment, and formation of cell debris observed in HUVECs and CAECs treated with LY83583, was blunted in EPCs, indicating stress tolerance of progenitor cells. Further study of apoptosis (one of the cell death mechanisms) using TUNEL assay demonstrated that EPCs were more resistant to apoptosis induced by serum depletion and LY83583 compared with mature endothelial cells (Figure 2D and 2E; representative fluorescent confocal micrographs shown in Figure IIA, available online at http://atvb.ahajournals.org). However, the apoptotic response of EPCs to cisplatin (a direct DNA damage agent)24 were similar to those in mature cells (Figure IIB). Matrigel tube formation assay (an in vitro angiogenic assay) showed that incubation of cells with 8 µmol/L LY83583 caused a severe inhibition of capillary tube formation by mature endothelial cells (HUVECs and CAECs), but not by EPCs (Figure 3 A and 3B).
|
|
Elevated Expression of MnSOD in EPCs
On Western blot, EPCs compared with HUVECs and CAECs displayed at baseline 3- to 4-fold higher protein levels of MnSOD, a mitochondrial antioxidant enzyme (Figure 4). This difference was consistently present in all tested passages (HUVECs, 2 to 9 passages; CAECs, 4 to 9 passages; EPCs, 4 to 8 passages; data not shown). This distinct expression profile translated into an increased enzymatic activity of MnSOD, which was found to be significantly higher in EPCs compared with HUVECs and CAECs (Figure 4). In contrast, the protein expressions and enzymatic activities of CuZnSOD or catalase were not significantly different between EPCs and mature endothelial cells. Moreover, TNF-, an inducer of MnSOD,25 produced significantly higher expression of this protective enzyme in EPCs compared with HUVECs, which persisted for at least 72 hours (Figure 5 A and 5B). TNF- (0.1 and 0.5 ng/mL, 24 hours) treatment did not significantly change the expression of CuZnSOD or catalase in EPCs and HUVECs (data not shown). Thus stress-tolerant EPCs display a distinct antioxidant profile characterized by an elevated constitutive expression of mitochondrial MnSOD.
|
|
Because induction of MnSOD by TNF- is nuclear factor 
B (NF-B)-dependent,26 we also examined 2 other proteins (VCAM-1 and COX-2) known to be induced by TNF- mediated by NF-B.27,28 As shown in Figure III (available online at http://atvb.ahajournals.org), TNF- induced VCAM-1 in both HUVECs and EPCs with significantly higher VCAM-1 expression in EPCs at concentrations of 0.01 and 0.1 ng/mL, whereas a higher VCAM-1 protein level was detected in HUVECs at concentrations of 0.5 ng/mL. In the same concentration range, TNF- stimulated COX-2 expression in HUVECs but not in EPCs (Figure III).
Mitochondrial Resistance of EPCs
EPCs, CAECs, and HUVECs demonstrated a similar intracellular distribution of mitochondria with a characteristic thread-like network appearance revealed by the mitochondrial-selective dye, Mito Tracker Red CM-H2Xros (29; Figure 6A). Under oxidative stress produced by 8 µmol/L LY83583 for 24 hours, in 92±4% of HUVECs and 87±3% of CAECs, mitochondrial staining showed disruption of the thread-like network with a significant punctate appearance (Figure 6A and 6B). In contrast, under the same stress, mitochondria in EPCs maintained the normal thread-shaped network with only a fraction of these cells (7±3%), demonstrating mitochondrial alteration and punctuation (Figure 6A and 6B). No punctate cell was found in non-LY83583-treated groups of HUVECs, CAECs, and EPCs (n=3 to 5). Live imaging of mitochondria with tetramethylrodamine methyl (TMRM) also demonstrated loss of normal mitochondrial architecture associated with generation of punctate-like mitochondria in the majority of LY83583-treated HUVECs but not EPCs (data not shown), indicative of selective mitochondrial tolerance in progenitor cells. At high resolution electron microscopy (Figure IVA, available online at http://atvb.ahajournals.org), the structural integrity of mitochondria in EPCs treated with LY83583 was maintained, whereas in HUVECs oxidative stress produced mitochondrial matrix swelling with organelle damage. Thus, EPCs demonstrate a higher mitochondrial tolerance toward oxidative stress.
|
To demonstrate that the cytotoxic effects of LY83583 are caused by increased formation of , and to further confirm that resistance of EPCs to oxidative stress is related to increase in MnSOD expression, CAECs were transduced with 50 MOI AdMnSOD or AdLacZ (as a control) before LY83583 treatment (8 µmol/L for 24 hours). Mitochondrial staining showed that mitochondria were resistant to oxidative damage by LY83583 in the majority of cells transduced with 50 MOI AdMnSOD but not in cells transduced with 50 MOI AdLacZ (Figure 6B). AdMnSOD or AdLacZ transduction alone did not cause significant mitochondrial damage (8±5% and 2±1% cells with punctate appearance in CAECs transduced with AdMnSOD and in CAECs transduced with AdLacZ, respectively; n=3). Live imaging of mitochondria by TMRM indicated that tranduction of HUVECs with 100 MOI AdMnSOD (but not with AdLacZ) protected mitochondria from damage by LY83583 (data not shown). Cell viability assayed by trypan blue exclusion demonstrated that 50 MOI AdMnSOD protected HUVECs from cytotoxicity of LY83583, whereas AdLacZ failed to show the protective effect (Figure IVB).
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
The results of this study demonstrate that EPCs are resistant to cytotoxic effects induced by LY83583, serum depletion, or TNF-
compared with mature endothelial cells (CAECs and HUVECs). This resistance is in part due to the ability of EPCs to preserve mitochondrial integrity under conditions of oxidative stress. High level of MnSOD expression and enzymatic activity in EPCs appear to be the major molecular mechanism underlying this phenomenon. In contrast, expression and function of CuZnSOD and catalase are similar in EPCs and CAECs. These results suggest that MnSOD is the most likely of the 3 antioxidant enzymes to be responsible for resistance of EPCs to oxidative stress. In the present study, the endothelial phenotype of EPCs was established by morphological, immunohistochemical, and biochemical criteria.6,7,9 EPCs expressed endothelial specific markers, whereas cytokines downregulated expression of eNOS in EPCs consistent with the reported ability of cytokines to decrease expression of eNOS in HUVECs.30 Interestingly, we observed a low basal level of eNOS protein expression in EPCs as compared with mature endothelial cells. Functional implications of low eNOS expression in EPCs are unclear and remain to be determined.
Oxidative stress plays an essential role in pathogenesis of vascular disease.13,31 In the present study, treatment with O·–2 generator LY83583 and conditions (TNF- and serum depletion) reported to induce ROS-mediated apoptosis in various systems22,23 caused cell death and abolished angiogenic capacity in HUVECs and CAECs but not in EPCs, suggesting that EPCs may have a higher antioxidant capacity than mature endothelial cells. Our study also showed that very high levels of oxidative stress could overwhelm the antioxidant defenses causing EPCs death, as demonstrated by treatment of EPCs with high concentration of LY83583 (12 µmol/L). Interestingly, EPCs were not resistant to DNA-damaging agent cisplatin. Even though increased production of ROS has been observed after treatment with DNA damaging agents such as cisplatin,32 current evidence suggests that this increase in ROS after DNA damage reflects caspase-mediated disruption of electron transport in mitochondria during the cell death process.33,34 The sensitivity of EPCs to cisplatin may be due to the possibility that the DNA damage and subsequent activation of proapoptosis pathways (rather than mitochondrial oxidative stress) are primarily responsible for cytochrome c release and apoptosis. Apparently, EPCs may not be resistant to all death-inducing stimuli.
Our studies demonstrate that EPCs had a higher expression and enzymatic activity of MnSOD compared with HUVECs and CAECs, although the protein expression and activity of CuZnSOD and catalase were similar between EPCs and mature endothelial cells. This finding suggests that MnSOD may play an important role in the resistance of EPCs to oxidative stress. Our results are consistent with the previously reported cytoprotective effect of MnSOD.35,36 Further support for this concept was provided by morphological analysis of mitochondria. In HUVECs, LY83583 induced swelling of mitochondria that was reflected in disruption of the normal intracellular mitochondrial network. Minimal morphological changes were detected in EPCs exposed to LY83583. Most importantly, overexpression of recombinant MnSOD in CAECs and HUVECs prevented LY83583-induced mitochondrial damage and cell death, strongly suggesting that MnSOD is an essential enzyme responsible for the maintenance of normal mitochondrial morphology in endothelial cells faced with high concentration of . Thus functional, biochemical, and morphological evidence supports the concept that EPCs may be able to survive severe oxidative stress in part because of high level of MnSOD expression and enzymatic activity.
The proinflammatory cytokine TNF- can cause both apoptosis and necrosis by activation of complex signal transduction pathways. These effects could be mediated by increased production of ROS in mitochondria37 or upregulation of NADPH oxidase.38 In contrast, TNF- has been shown to protect cells against ROS by increasing expression and activity of MnSOD.25 In this study, we provide the first evidence that TNF- causes a concentration-dependent upregulation of MnSOD protein expression in EPCs. This effect was much more pronounced than the corresponding effect of TNF- in HUVECs, suggesting that EPCs may be able to withstand more severe oxidative stress than adult endothelium under pathological conditions associated with increasing tissue concentration of cytokines.39 The mechanisms underlying high induction of MnSOD in EPCs are not clear. TNF--induced expressions of MnSOD, VCAM-1, and COX-2 are NF-B-dependent.26–28 However, regulation of MnSOD by TNF- in EPCs apparently is different from COX-2 and VCAM-1. In EPCs, we were not able to induce expression of COX-2 protein by TNF-. Furthermore, the induction of VCAM-1 by TNF- was stronger in EPCs in the presence of low concentrations of TNF-. In contrast, higher concentrations of TNF- induced higher expression of VCAM-1 in HUVECs as compared with EPCs. Thus it appears that in EPCs, TNF- selectively upregulates expression of MnSOD. Mechanisms responsible for this selectivity remain to be investigated.
Despite the limitations inherent in culturing of endothelial cells of different origin, the results of this study demonstrate that human EPCs isolated from circulating blood are resistant to cytotoxic effects induced by LY83583, TNF-, and serum depletion. Complex molecular mechanisms could be responsible for this phenomenon. The results of our experiments point to MnSOD as an important component of EPC resistance to oxidative stress. We speculate that the elevated antioxidant capacity is designed to enable EPCs to survive in a high oxidative stress environment during ischemia-induced angiogenesis, ischemia-reperfusion, and/or inflammation.
|
Acknowledgments |
|---|
This work was supported in part by National Heart, Lung, and Blood Institute grants HL-53524, HL-58080, and HL-066958, by the American Heart Association Bugher Award for Investigation of Stroke, and the Mayo Foundation. The authors thank Dr Scott Kaufman and Dr Xue Meng for helpful discussions during revision of this manuscript. Secretarial assistance of Janet Beckman is gratefully acknowledged.
Received April 2, 2004; accepted July 26, 2004.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
1. Iba O, Matsubara H, Nozawa Y, Fujiyama S, Amano K, Mori Y, Kojima H, Iwasaka T. Angiogenesis by implantation of peripheral blood mononuclear cells and platelets into ischemic limbs. Circulation. 2002; 106: 2019–2025.
2. Asahara T, Masuda H, Takahashi T, Kalka C, Pastore C, Silver M, Kearne M, Magner M, Isner J. M. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Cir Res. 1999; 85: 221–228.
3. Otani A, Kinder K, Ewalt K, Otero FJ, Schimmel P, Friedlander M. Bone marrow-derived stem cells target retinal astrocytes and can promote or inhibit retinal angiogenesis. Nat Med. 2002; 8: 1004–1010.[CrossRef][Medline] [Order article via Infotrieve]
4. Assmus B, Schachinger V, Teupe C, Britten M, Lehamann R, Dobert N, Grunwald F, Aicher A, Urbich C, Martin H, Hoelzer D, Dimmeler S, Zeiher AM. Transplantation of progenitor cells and regeneration enhancement in acute myocardial infarction. Circulation. 2002; 106: 3009–3017.
5. Griese DP, Ehsan A, Melo LG, Kong D, Zhang L, Mann MJ, Pratt RE, Mulligan RC, Dzau VJ. Isolation and transplantation of autologous circulating endothelial cells into denuded vessels and prosthetic grafts: implications for cell-based vascular therapy. Circulation. 2003; 108: 2710–2715.
6. Asahara T, Murohara T, Sullivan A, Silver M, Van der Zee R, Li T, Witzenbichler B, Schatteman G, Isner JM. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997; 275: 964–967.
7. Lin Y, Weisdorf DJ, Solovey A, Hebbel RP. Origins of circulating endothelial cells and endothelial outgrowth from blood. J Clin Invest. 2000; 105: 71–77.[Medline] [Order article via Infotrieve]
8. Majka SM, Jackson KA, Kienstra KA. Majesky MW, Goodell MA, Hirschi KK. Distinct progenitor populations in skeletal muscle are bone marrow derived and exhibit different cell fates during vascular regeneration. J Clin Invest. 2003; 111: 71–79.[CrossRef][Medline] [Order article via Infotrieve]
9. Simper D, Stalboerger PG, Panetta CJ, Wang S, Caplice NM. Smooth muscle progenitor cells in human blood. Circulation. 2002; 106: 1199–1204.
10. Kawamoto A, Gwon HC, Iwaguro H, Yamaguchi JI, Uchida S, Masuda H, Silver M, Ma H, Kearney M, Isner JM, Asahara T. Therapeutic potential of ex vivo expanded endothelial progenitor cells for myocardial ischemia. Circulation. 1998; 103: 634–637.
11. Kalka C, Masuda H, Takahashi T, Kalka-Moll WM, Silver M, Kearney M, Li T, Isner JM, Asahara T. Transplantation of ex vivo expanded endothelial progenitor cells for therapeutic neovascularization. Proc Natl Acad Sci U S A. 2000; 97: 3422–3427.
12. Tateishi-Yuyama E, Matsubara H, Murohara T, Ikeda U, Shintani S, Masaki H, Amano K, Kishimoto Y, Yoshimoto K, Akashi H, Shimada K, Iwasaka T, Imaizumi T. Therapeutic angiogenesis for patients with limb ischaemia by autologous transplantation of bone-marrow cells: a pilot study and a randomised controlled trial. Lancet. 2002; 360: 427–435.[CrossRef][Medline] [Order article via Infotrieve]
13. Cai H, Harrison DG. Endothelial dysfunction in cardiovascular disease, the role of oxidant stress. Cir Res. 2000; 87: 840–844.
14. Oberley LW, Buettner GR. Role of superoxide dismutase in cancer: a review. Cancer Res. 1979; 39: 1141–1149.
15. Schmid I, Uittenbogaart CH, Giorgi JV. A gentle fixation and permeabilization method for combined cell surface and intracellular staining with improved precision in DNA quantification. Cytometry. 1991; 12: 279–285.[CrossRef][Medline] [Order article via Infotrieve]
16. He T, Weintraub NL, Goswami PC, Chatterjee P, Flaherty DM, Domann FE. Oberley LW. Redox factor-1 contributes to the regulation of progression from G0/G1 to S by PDGF in vascular smooth muscle cells. Am J Physiol Heart Cir Physiol. 2003; 285: H804–H812.
17. Li S, Yan T, Yang JQ, Oberley TD, Oberley LW. The role of cellular glutathione peroxidase redox regulation in the suppression of tumor cell growth by manganese superoxide dismutase. Cancer Res. 2000; 60: 3927–3939.
18. Dzeja PP, Bortolon R, Perez-Terzic C, Holmuhamedov EL, Terzic A. Energetic communication between mitochondria and nucleus directed by catalyzed phosphotransfer. Proc Natl Acad Sci U S A. 2002; 99: 10156–10161.
19. Beauchamp C, Fridovich I. Superoxide dismutase: improved assays and an assay applicable to acrylamide gels. Anal Biochem. 1971; 44: 276–287.[CrossRef][Medline] [Order article via Infotrieve]
20. Sun Y, Elwell JH, Oberley LW. A simultaneous visualization of the antioxidant enzymes glutathione peroxidase and catalase on polyacrylamide gels. Free Rad Res Commun. 1988; 5: 67–75.[Medline] [Order article via Infotrieve]
21. Peterson TE, Poppa V, Ueba H, Wu A, Yan C, Berk BC. Opposing effects of reactive oxygen species and cholesterol on endothelial nitric oxide synthase and endothelial cell caveolae. Cir Res. 1999; 85: 29–37.
22. Manna SK, Mukhopadhyay A, Aggarwal BB. Resveratrol suppresses TNF-induced activation of nuclear transcription factors NF-kappa B, activator protein-1, and apoptosis: potential role of reactive oxygen intermediates and lipid peroxidation. J Immunol. 2000; 164: 6509–6519.
23. Andoh T, Chock PB, Chiueh CC. The roles of thioredoxin in protection against oxidative stress-induced apoptosis in SH-SY5Y cells. J Biol Chem. 2002; 277: 9655–9660.
24. Seluanov A, Gorbunova V, Falcovitz A, Sigal A, Milyavsky M, Zurer I, Shohat G, Goldfinger N, Rotter V. Change of the death pathway in senescent human fibroblasts in response to DNA damage is caused by an inability to stabilize p53. Mol Cell Biol. 2001; 21: 1552–1564.
25. Wong GH, Elwell JH, Oberley LW, Goeddel DV. Manganous superoxide dismutase is essential for cellular resistance to cytotoxicity of tumor necrosis factor. Cell. 1989; 58: 923–931.[CrossRef][Medline] [Order article via Infotrieve]
26. Guo Z, Boekhoudt GH, Boss JM. Role of the intronic enhancer in tumor necrosis factor-mediated induction of manganous superoxide dismutase. J Biol Chem. 2003; 278: 23570–23578.
27. Neish AS, Khachigian LM, Park A, Baichwal VR, Collins T. Sp1 is a component of the cytokine-inducible enhancer in the promoter of vascular cell adhesion molecule-1. J Biol Chem. 1995; 70: 28903–28909.
28. Huang WC, Chen JJ, Inoue H, Chen CC. Tyrosine phosphorylation of I-kappa B kinase alpha/beta by protein kinase C-dependent c-Src activation is involved in TNF--induced cyclooxygenase-2 expression. J Immunol. 2003; 170: 4767–4775.
29. Degli Esposti M. Measuring mitochondrial reactive oxygen species. Methods. 2002; 26: 335–340.[CrossRef][Medline] [Order article via Infotrieve]
30. Rosenkranz-Weiss P, Sessa WC, Milstien S, Kaufman S, Watson CA, Pober JS. Regulation of nitric oxide synthesis by proinflammatory cytokines in human umbilical vein endothelial cells. Elevations in tetrahydrobiopterin levels enhance endothelial nitric oxide synthase specific activity. J Clin Invest. 1994; 93: 2236–2243.
31. Harrison D, Griendling KK, Landmesser U, Horning B, Drexler H. Role of oxidative stress in atherosclerosis. Am J Cardiol. 2003; 91: 7A–11A.[CrossRef][Medline] [Order article via Infotrieve]
32. Baek SM, Kwon CH, Kim JH, Woo JS, Jung JS, Kim YK. Differential roles of hydrogen peroxide and hydroxyl radical in cisplatin-induced cell death in renal proximal tubular epithelial cells. J Lab Clin Med. 2003; 142: 178–186.[CrossRef][Medline] [Order article via Infotrieve]
33. Bossy-Wetzel E, Newmeyer DD, Green DR. Mitochondrial cytochrome c release in apoptosis occurs upstream of DEVD-specific caspase activation and independently of mitochondrial transmembrane depolarization. EMBO J. 1998; 17: 37–49.[CrossRef][Medline] [Order article via Infotrieve]
34. Ricci JE, Muñoz-Pinedo C, Fitzgerald P, Bailly-Maitre B, Perkins GA, Yadava N, Scheffler IE, Ellisman MH, Green DR. Disruption of Mitochondrial Function during Apoptosis Is Mediated by Caspase Cleavage of the p75 Subunit of Complex I of the Electron Transport Chain. Cell. 2004; 117: 773–786.[CrossRef][Medline] [Order article via Infotrieve]
35. Li JJ, Oberley LW. Overexpression of manganese-containing superoxide dismutase confers resistance to the cytotoxicity of tumor necrosis factor and/or hyperthermia. Cancer Res. 1997; 57: 1991–1998.
36. Melov S, Schneider JA, Day BJ, Hinerfeld D, Coskun P, Mirra SS, Crapo JD, Wallace DC. A novel neurological phenotype in mice lacking mitochondrial manganese superoxide dismutase. Nat Gen. 1998; 18: 159–163.[CrossRef][Medline] [Order article via Infotrieve]
37. Goossens V, Grooten De Vos K, Fiers WJ. Direct evidence for tumor necrosis factor-induced mitochondrial reactive oxygen intermediates and their involvement in cytotoxicity. Proc Natl Acad Sci U S A. 1995; 92: 8115–8119.
38. Ungvari Z, Csiszar A, Edwards JG, Kaminski PM, Wolin MS, Kaley G, Koller A. Increased superoxide production in coronary arteries in hyperhomocysteinemia: role of tumor necrosis factor-, NAD(P)H oxidase, and inducible nitric oxide synthase. Arterioscler, Thromb Vasc Biol. 2003; 23: 418–424.
39. Levine B, Kalman J, Mayer L, Fillit H, Packer M. Elevated circulating levels of tumor necrosis factor in severe chronic heart failure. N Engl J Med. 1990; 323: 236–241.[Abstract]
Related Article:
Arterioscler. Thromb. Vasc. Biol. 2004 24: 1977-1979.
This article has been cited by other articles:
 |
|
 |
|
  M. Thurau, G. Marquardt, N. Gonin-Laurent, K. Weinlander, E. Naschberger, R. Jochmann, K. R. Alkharsah, T. F. Schulz, M. Thome, F. Neipel, et al. Viral Inhibitor of Apoptosis vFLIP/K13 Protects Endothelial Cells against Superoxide-Induced Cell Death J. Virol., January 15, 2009; 83(2): 598 - 611. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. L. Urish, J. B. Vella, M. Okada, B. M. Deasy, K. Tobita, B. B. Keller, B. Cao, J. D. Piganelli, and J. Huard Antioxidant Levels Represent a Major Determinant in the Regenerative Capacity of Muscle Stem Cells Mol. Biol. Cell, January 1, 2009; 20(1): 509 - 520. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. He, T. Lu, L. V. d'Uscio, C.-F. Lam, H.-C. Lee, and Z. S. Katusic Angiogenic Function of Prostacyclin Biosynthesis in Human Endothelial Progenitor Cells Circ. Res., July 3, 2008; 103(1): 80 - 88. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Dimmeler and A. Leri Aging and Disease as Modifiers of Efficacy of Cell Therapy Circ. Res., June 6, 2008; 102(11): 1319 - 1330. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Spinetti, N. Kraenkel, C. Emanueli, and P. Madeddu Diabetes and vessel wall remodelling: from mechanistic insights to regenerative therapies Cardiovasc Res, May 1, 2008; 78(2): 265 - 273. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A.-L. Levonen, E. Vahakangas, J. K. Koponen, and S. Yla-Herttuala Antioxidant Gene Therapy for Cardiovascular Disease: Current Status and Future Perspectives Circulation, April 22, 2008; 117(16): 2142 - 2150. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Dulak, J. Deshane, A. Jozkowicz, and A. Agarwal Heme Oxygenase-1 and Carbon Monoxide in Vascular Pathobiology: Focus on Angiogenesis Circulation, January 15, 2008; 117(2): 231 - 241. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. A. Sorrentino, F. H. Bahlmann, C. Besler, M. Muller, S. Schulz, N. Kirchhoff, C. Doerries, T. Horvath, A. Limbourg, F. Limbourg, et al. Oxidant Stress Impairs In Vivo Reendothelialization Capacity of Endothelial Progenitor Cells From Patients With Type 2 Diabetes Mellitus: Restoration by the Peroxisome Proliferator-Activated Receptor-{gamma} Agonist Rosiglitazone Circulation, July 10, 2007; 116(2): 163 - 173. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. K. Kissel, R. Lehmann, B. Assmus, A. Aicher, J. Honold, U. Fischer-Rasokat, C. Heeschen, I. Spyridopoulos, S. Dimmeler, and A. M. Zeiher Selective Functional Exhaustion of Hematopoietic Progenitor Cells in the Bone Marrow of Patients With Postinfarction Heart Failure J. Am. Coll. Cardiol., June 19, 2007; 49(24): 2341 - 2349. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y.-H. Chen, S.-J. Lin, F.-Y. Lin, T.-C. Wu, C.-R. Tsao, P.-H. Huang, P.-L. Liu, Y.-L. Chen, and J.-W. Chen High Glucose Impairs Early and Late Endothelial Progenitor Cells by Modifying Nitric Oxide-Related but Not Oxidative Stress-Mediated Mechanisms Diabetes, June 1, 2007; 56(6): 1559 - 1568. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Kubo, T.-S. Li, R. Suzuki, M. Ohshima, S.-L. Qin, and K. Hamano Short-term pretreatment with low-dose hydrogen peroxide enhances the efficacy of bone marrow cells for therapeutic angiogenesis Am J Physiol Heart Circ Physiol, June 1, 2007; 292(6): H2582 - H2588. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Lamping Endothelial Progenitor Cells: Sowing the Seeds for Vascular Repair Circ. Res., May 11, 2007; 100(9): 1243 - 1245. [Full Text] [PDF]
|
|
|
|
|
|
A. V. R. Santhanam, L. A. Smith, T. He, K. A. Nath, and Z. S. Katusic Endothelial Progenitor Cells Stimulate Cerebrovascular Production of Prostacyclin By Paracrine Activation of Cyclooxygenase-2 Circ. Res., May 11, 2007; 100(9): 1379 - 1388. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. A. Ingram, T. R. Krier, L. E. Mead, C. McGuire, D. N. Prater, J. Bhavsar, M. R. Saadatzadeh, K. Bijangi-Vishehsaraei, F. Li, M. C. Yoder, et al. Clonogenic Endothelial Progenitor Cells Are Sensitive to Oxidative Stress Stem Cells, February 1, 2007; 25(2): 297 - 304. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Lu, T. He, Z. S. Katusic, and H.-C. Lee Molecular Mechanisms Mediating Inhibition of Human Large Conductance Ca2+-Activated K+ Channels by High Glucose Circ. Res., September 15, 2006; 99(6): 607 - 616. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Haendeler and S. Dimmeler Inseparably Tied: Functional and Antioxidative Capacity of Endothelial Progenitor Cells Circ. Res., February 3, 2006; 98(2): 157 - 158. [Full Text] [PDF]
|
|
|
|
|
|
G. Galasso, S. Schiekofer, K. Sato, R. Shibata, D. E. Handy, N. Ouchi, J. A. Leopold, J. Loscalzo, and K. Walsh Impaired Angiogenesis in Glutathione Peroxidase-1-Deficient Mice Is Associated With Endothelial Progenitor Cell Dysfunction Circ. Res., February 3, 2006; 98(2): 254 - 261. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. M. Smadja, I. Bieche, G. Uzan, H. Bompais, L. Muller, C. Boisson-Vidal, M. Vidaud, M. Aiach, and P. Gaussem PAR-1 Activation on Human Late Endothelial Progenitor Cells Enhances Angiogenesis In Vitro With Upregulation of the SDF-1/CXCR4 System Arterioscler. Thromb. Vasc. Biol., November 1, 2005; 25(11): 2321 - 2327. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C.-H. Yoon, J. Hur, K.-W. Park, J.-H. Kim, C.-S. Lee, I.-Y. Oh, T.-Y. Kim, H.-J. Cho, H.-J. Kang, I.-H. Chae, et al. Synergistic Neovascularization by Mixed Transplantation of Early Endothelial Progenitor Cells and Late Outgrowth Endothelial Cells: The Role of Angiogenic Cytokines and Matrix Metalloproteinases Circulation, September 13, 2005; 112(11): 1618 - 1627. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. He, T. E. Peterson, and Z. S. Katusic Paracrine mitogenic effect of human endothelial progenitor cells: role of interleukin-8 Am J Physiol Heart Circ Physiol, August 1, 2005; 289(2): H968 - H972. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
O. Awad, C. Jiao, N. Ma, M. Dunnwald, and G. C. Schatteman Obese Diabetic Mouse Environment Differentially Affects Primitive and Monocytic Endothelial Cell Progenitors Stem Cells, April 1, 2005; 23(4): 575 - 583. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. Rossig, C. Urbich, and S. Dimmeler Endothelial Progenitor Cells at Work: Not Mature Yet, but Already Stress-Resistant Arterioscler. Thromb. Vasc. Biol., November 1, 2004; 24(11): 1977 - 1979. [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||