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BioMed Research International
Volume 2013 (2013), Article ID 190486, 11 pages
Vasculogenic Cytokines in Wound Healing
Department of Surgery, Oregon Health and Science University, 3181 SW Sam Jackson Park Road, Portland, OR 97239, USA
Received 1 October 2012; Revised 8 January 2013; Accepted 23 January 2013
Academic Editor: Jorge Berlanga Acosta
Copyright © 2013 Victor W. Wong and Jeffrey D. Crawford. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Chronic wounds represent a growing healthcare burden that particularly afflicts aged, diabetic, vasculopathic, and obese patients. Studies have shown that nonhealing wounds are characterized by dysregulated cytokine networks that impair blood vessel formation. Two distinct forms of neovascularization have been described: vasculogenesis (driven by bone-marrow-derived circulating endothelial progenitor cells) and angiogenesis (local endothelial cell sprouting from existing vasculature). Researchers have traditionally focused on angiogenesis but defects in vasculogenesis are increasingly recognized to impact diseases including wound healing. A more comprehensive understanding of vasculogenic cytokine networks may facilitate the development of novel strategies to treat recalcitrant wounds. Further, the clinical success of endothelial progenitor cell-based therapies will depend not only on the delivery of the cells themselves but also on the appropriate cytokine milieu to promote tissue regeneration. This paper will highlight major cytokines involved in vasculogenesis within the context of cutaneous wound healing.
It is estimated that diabetic and cardiovascular complications will account for 9 trillion in US healthcare costs over the next thirty years . These complications are often associated with impaired blood vessel growth in response to tissue hypoxia and ischemia. Chronic nonhealing wounds represent an important public health problem as populations prone to impaired wound healing continue to grow (e.g., diabetics, elderly, and obese) . The estimated healthcare cost of diabetic foot ulcers alone has been estimated at 45,000 per patient [3, 4]. Thus, strategies to augment the neovascularization response to injury may dramatically improve the quality of life for these patients and significantly reduce the global biomedical burden [1, 5, 6].
Regulation of blood vessel development in response to tissue injury or ischemia is critical for maintenance of healthy tissues . A robust vascular response to deliver immune cells and metabolic substrates is important for cutaneous wound healing . In addition, coordinated neovascularization programs are essential for normal organ development during embryogenesis . Conversely, dysregulated signaling can promote tumor growth and metastasis [10, 11]. A better understanding of blood vessel formation in both health and disease states may result in more effective therapies for a wide range of diseases.
During embryogenesis, mesoderm-derived angioblasts organize to form blood vessels via vasculogenesis . It was initially believed that all subsequent blood vessel growth occurred through sprouting of preexisting endothelial cells via angiogenesis . However, it is now known that the vascular programming present during embryonic development is recapitulated in various postnatal states during a process known as adult vasculogenesis  (Figure 1). Vasculogenesis plays a critical role in maintaining tissue homeostasis throughout the body . Disruption of these pathways can sustain pathogenic processes (e.g., in skin, heart, kidney, and brain) that are only starting to be appreciated on a molecular level. The remainder of this paper refers to postnatal vasculogenesis and focuses on major vasculogenic cytokines in the clinical context of wound healing.
2. Endothelial Precursor and Other Provasculogenic Cells
Endothelial precursor cells (EPCs) are bone-marrow-derived progenitor cells that participate in vasculogenesis and were first identified by Asahara et al. . These cells are recruited to sites of ischemia and divide to form syncytial masses which tubularize and canalize to form a patent vascular network . Although the molecular identification of EPCs remains a topic of debate, studies suggest that two functionally distinct subpopulations exist based on in vitro isolation techniques: early outgrowth EPCs and late outgrowth EPCs [18, 19]. Specifically, early outgrowth EPCs appear to function in a paracrine role in promoting neovascularization whereas late outgrowth EPCs directly differentiate into endothelial tubules . Transcriptional and proteomic profiling of these populations suggests that early outgrowth EPCs may be of monocytic origin and restricted in their ability to promote neovascularization clinically .
EPCs have also been characterized based on their surface expression profiles . In human studies, combinations of surface markers used to identify EPCs often include CD34+, CD133+, and VEGFR-2+. In mice, common EPC surface markers include Sca-1+, Lin-, Flk-1+, and cKit+. It is important to note that none of the markers used are specific for EPCs. Regardless of how they are classified, a common feature of EPCs is their ability to mobilize and home to injured areas and promote vessel formation . Various signaling molecules are highly implicated in this process and include transforming growth factor beta (TGF) and matrix metalloproteinase-9 (MMP-9) .
EPCs are thought to mobilize from the bone-marrow or other tissues and home to areas of endothelial damage via adhesion molecules. The secreted proteases cathepsin L and MMP2 regulate the transmigration of EPCs, which subsequently mature and differentiate towards the endothelial lineage [24, 25]. Nitric oxide signaling and reactive oxygen species have also been implicated in EPC activity, potentially affecting their colony-forming potential and ability to counteract ischemic stress . Nitric oxide pathways have even been linked to the ability of hormonal estrogens to promote EPC proliferation and mobilization .
Another cell population intimately involved in vascular morphogenesis is the pericyte, a supportive stromal-like cell that retains the pluripotency of mesenchymal stem cells (MSCs) . They reside at the interface between endothelial cells and the surrounding tissue, producing proangiogenic signals that regulate endothelial cell differentiation and growth . Through both direct physical interaction and paracrine signaling, endothelial cells and pericytes engage in complex crosstalk that is essential for normal adult vasculogenesis  (Figure 2). Fibroblasts have also been shown to facilitate EPC migration, branching, and sprouting in collagen matrices in vitro, potentially via cytokine signaling . Finally, platelets are synergistically involved in vasculogenesis, elaborating potent cytokines that regulate the recruitment and differentiation of EPCs . Diverse cell types are clearly involved in the formation of new blood vessels and the cytokine networks through which they communicate play a critical role in the tissue response to injury.
3. Major Vasculogenic Cytokines (Table 1)
Vascular endothelial growth factors (VEGF) are a family of cytokines important in both embryonic and post-natal vascular development . They play a crucial role in endothelial cell motility, proliferation, and survival . This wide range of effects is mediated in part by the multiple VEGF subtypes and the associated family of VEGF receptor (VEGFR) protein tyrosine kinases. Five human VEGF isoforms (A, B, C, D, and placental growth factor-PlGF) are produced by differential splicing of VEGF mRNA. VEGF-A is involved in vascular growth, lymphatic development, and vascular malformations . The role of VEGF-B in vascular development is poorly understood but may be associated with blocking apoptosis . VEGF-C and VEGF-D are involved in lymphangiogenesis, and PIGF appears to regulate angiogenesis, wound repair, and inflammation .
VEGF-A has been shown to promote adult vasculogenesis via bone-marrow-derived EPC mobilization , a process that acts via VEGFR1 and VEGFR2 in a tumor model . In a small animal model of soft tissue ischemia, VEGF levels and circulating VEGFR2+ cells were increased following injury, findings that correlated with migration of EPC populations to ischemic tissue . Furthermore, it has been shown that topical VEGF delivery can improve diabetic wound healing in a murine model through local upregulation of angiogenic cytokines and recruitment of bone-marrow-derived vasculogenic cells .
VEGF has also been shown to regulate the expression of endothelial cell surface proteins known as integrins that link cells with the extracellular matrix. Integrins, which comprise a family of transmembrane heterodimeric proteins, play a major role in controlling EPC mobilization and homing to areas of tissue injury and ischemia . Specifically, integrin has been shown to promote VEGF-induced differentiation of EPCs in vitro, highlighting the importance of both structural and cytokine signals in regulating EPC activity following injury.
A key mechanism that regulates VEGF expression is the hypoxia-inducible factor-1 (HIF-1) pathway. HIF-1 is a transcription factor that exists as a dimeric complex consisting of a cytoplasmic subunit and a nuclear subunit . In the setting of hypoxia, HIF-1 (which is degraded under normoxic conditions) translocates into the nucleus to complex with HIF-1, initiating the transcription of neovascularization genes including VEGF. Studies have demonstrated that impaired HIF-1 binding to its coactivator p300 may underlie diabetic impairments in wound healing . Thus, strategies to stabilize HIF-1 may enhance EPC mobilization and function  and have been shown to improve cutaneous wound healing in diabetic mice [42, 44].
Stromal cell-derived factor-1 (SDF-1) is a chemokine which plays a crucial role in EPC and hematopoietic stem cell (HSC) trafficking through the circulation . SDF-1 binds exclusively to the chemokine receptor CXCR4, which is expressed by circulating cells and regulates their recruitment from bone-marrow . In addition, SDF-1 mediates the activation of circulating stem cells during embryonic organogenesis and vascular development , suggesting that it may serve similar functions in post-natal neovascularization. Dysfunctional SDF-1 pathways have been highly implicated in aged and diabetic wound healing in preclinical models , underscoring the importance of chemokine-mediated signaling networks in normal wound healing.
Researchers have examined the role of SDF-1 in peripheral vasculogenesis and tissue repair. It has been demonstrated that SDF-1 gene expression in EPCs is regulated by the transcription factor HIF-1 and that cutaneous tissues express SDF-1 in response to hypoxia . In addition, blockage of either SDF-1 or its receptor CXCR4 can prevent stem cell recruitment to ischemic tissues. Local delivery of SDF-1 into ischemic muscle has been shown to enhance vasculogenesis via EPC recruitment , highlighting its potential as a therapeutic chemokine. Additionally, plasmid gene transfer of SDF-1 has been demonstrated to augment neovascularization through VEGF . In the setting of diabetic wound healing, administration of SDF-1 is capable of reversing the impairment in EPC homing to injured tissue . Other forms of tissue injury have also been shown to activate SDF-1. In a mouse burn wound model, researchers have characterized SDF-1 expression in the healing margin of burn wounds . Ionizing radiation injury also appears to stimulate vessel formation via SDF-1, however in a HIF-independent manner . Together, these studies collectively highlight the importance of SDF-1 in regulating wound vasculogenesis and suggest a role for chemokines in the treatment of chronic wounds.
The platelet-derived growth factor (PDGF) family of ligands and receptors is closely related to VEGF and may have evolved from a common gene . The family of four ligands (PDGF-A, PDGF-B, PDGF-C, and PDGF-D) assemble intracellularly and undergo transcriptional and posttranslational modifications. Specifically, the homodimer PDGF-BB recruits perivascular cells during vasculogenesis, possibly through the generation of reactive oxygen species and subsequent activation of extracellular-regulated kinase 1, 2 (ERK 1, 2) . Endothelial-derived PDGF-BB also induces progenitor cell migration and expansion during vascular development  and is critical during vascular bed formation by mesangial progenitor cells . VEGF ligands can also bind and activate PDGF pathways, a process important during MSC-associated vasculogenesis .
Researchers have exploited PDGF pathways to control neovascularization in various animal models. For example, nanofibrous scaffolds incorporated with PDGF have been shown to activate cytokine signaling and improve angiogenesis during wound repair in rats . Additionally, a constitutively activating mutation of the PDGF receptor was introduced into embryonic stem cells and shown to enhance vascular development both in vivo and in vitro, potentially through VEGF pathways . Furthermore, a PDGF-receptor antagonist has been shown to inhibit human tumor growth in a rat model, an effect that was augmented using anti-VEGF antibody .
In human studies, neovessels in revascularized wounds exhibit strong PDGF receptor staining . These findings are consistent with data demonstrating that PDGF is a primary mediator of vessel maturation . PDGF-dependent pathways are thought to drive angiogenic sprouting and vessel enlargement via vascular cell migration and proliferation . In fact, PDGF was the first growth factor to be approved by the United States Food and Drug Administration for the clinical treatment of ulcers . Taken together, these studies indicate that PDGF signaling is closely associated with VEGF pathways and is important during both developmental and adult vasculogenesis.
The fibroblast growth factor (FGF) family of cytokines displays diverse functional properties that are important in multiple aspects of wound repair including vasculogenesis . Although FGF and VEGF differentially activate genes and stimulate the development of different vessel types, FGF appears to induce a vasculogenic response that is highly dependent on VEGF . In myocardial tissues, FGF-2 has been shown to augment angiogenesis and vascular remodeling in response to ischemic injury . FGF-1 has been used to induce neovascularization in both an omentum model and a vascular pedicle model in rats [68, 69], suggesting that FGF-based strategies may be effective in promoting blood vessel formation in complex tissue constructs. Recently, researchers demonstrated improved neovascularization in a murine hindlimb ischemia model using an FGF-based hydrogel delivery system .
EPCs express receptors for FGF and a subpopulation of CD34-expressing HSCs that specifically expresses FGFR-1 has been shown to differentiate into endothelial cells in vitro . FGF-1 has been shown to regulate the proliferation and differentiation of EPC-like mesenchymal cells , suggesting it may control neovascularization mediated by endothelial-stromal cell interactions. FGF also appears to function via autocrine and paracrine mechanisms in endothelial cells  and may play a role in tumor angiogenesis and invasiveness . Recently, researchers topically applied EPCs to diabetic wounds in mice and detected increased local expression of FGF and VEGF which corresponded with improved wound healing and vascularization , supporting a key role for these cytokines in vasculogenesis during soft tissue repair.
Granulocyte macrophage colony stimulating factor (GM-CSF) is a potent cytokine that stimulates the mobilization of hematopoietic progenitor/myeloid cells and nonhematopoietic cells (e.g., bone-marrow MSCs) . During wound healing, multiple cell types including keratinocytes, fibroblasts, macrophages, endothelial cells, dendritic cells, and lymphocytes secrete GM-CSF. It has been shown to directly promote reepithelialization and induce secondary cytokine secretion from various wound healing cells . Clinical trials have demonstrated the efficacy of topically applied recombinant human GM-CSF for deep partial thickness burn wounds, highlighting the importance of this cytokine in human wound repair .
GM-CSF is related to interleukins (IL)-3 and IL-5 and plays diverse roles in homeostasis and disease . Its role in angiogenesis is partly mediated by monocytes and VEGF-associated pathways . In human endothelial cells, GM-CSF activates intracellular phosphatidyl-inositol-3-kinase and Jak/Stat signaling during vascular tubule formation in vitro. Interestingly, immune defense pathways have been associated with GM-CSF-stimulated angiogenesis and may represent an integrated mechanism for tissue defense and regeneration following injury .
GM-CSF has also been closely linked to vasculogenic processes. It has been shown to stimulate EPC tubule formation, proliferation, migration, and viability in a dose- and time-dependent manner, effects which were mediated in part by ERK signaling and upregulation of VEGF and integrin 2 . GM-CSF has also been shown to enhance EPC recruitment and vasculogenesis in murine and rabbit hindlimb ischemia models , potentially via direct activation of endothelial cells during neovascularization . GM-CSF pathways have also been implicated in tumor vasculogenesis , indicating that it regulates blood vessel formation in both health and disease states.
Sphingosine-1-phosphate (S1P) is a sphingolipid metabolite found in high concentrations in blood and implicated in vascular development. It is secreted most prominently by platelets, suggesting that it may have an important role in tissue repair. Further, it has been shown to act via distinct receptor pathways to regulate keratinocyte and fibroblast chemotaxis, processes that are critical for normal wound healing [85, 86]. As proof of concept, subcutaneous injections of S1P were able to significantly improve diabetic wound healing and neovascularization in rodent models .
Gradients in S1P levels are known to mediate the migration of endothelial cells, potentially through a recently identified S1P transporter (SPNS2) . S1P is thought to stabilize vasculature in part through regulation of VEGF pathways and cadherins junctions, processes potentially altered by blood flow mechanotransduction signaling . Cadherin and S1P pathways have also been linked to vascular development in a zebrafish model , suggesting a key role for S1P in maintaining vascular integrity.
During embryonic vasculogenesis in mice, S1P has been demonstrated to promote migration of angioblasts and endothelial cells . S1P pathways have also been implicated in blood vessel development. For example, mice lacking the receptor for S1P displayed immature vessels that lacked pericytes and smooth muscle elements . Furthermore, other growth factors such as PDGF may act through sphingolipid signaling to promote cellular motility during blood vessel development , highlighting the functional diversity of this signaling pathway in vascular biology.
3.7. MMP-9 and Other Proteases
MMP-9 is a soluble extracellular protease that plays diverse roles in wound repair. Paradoxically, high levels of MMP-9 have been implicated in chronic nonhealing wounds as well as scarless wound repair in athymic mice [94, 95]. However, mice that lack MMP-9 also exhibit delayed wound healing with disordered collagen remodeling, suggesting that tight regulation of this protease is critical for normal cutaneous repair and remodeling . Recent studies suggest that keratinocyte secretion of MMP-9 may be crucial to maintain normal basement membrane and matrix integrity .
The formation of new blood vessels involves not only cellular motility, growth, and sprouting, but also dynamic interactions with the endothelial basement membrane. Integrin-laminin interactions have been shown to regulate vessel branching  while recruited pericytes play an active role in vascular morphogenesis . Matrix remodeling pathways are also highly involved in neovascularization, controlling neovessel growth, maturation, and regression during tissue repair . Specific proteases such as MMP-9 and their inhibitors regulate major aspects of extracellular matrix turnover and degradation during vascular remodeling .
In addition to local effects at the injury site, MMP-9 has been shown to recruit EPCs from the bone-marrow  and can induce matrix release of vasculogenic cytokines including VEGF and TGF [103, 104]. Studies using MMP-9 knockout mice have demonstrated that MMP-9 is essential for tumor vascularization  and augments EPC mobilization and migration in a hindlimb ischemia model . Additionally, stem-cell-activating cytokines may be released from the extracellular matrix by MMPs , further potentiating the neovascularization process.
Other proteases implicated in vascular formation include membrane-type MMPs (MT-MMPs) that act on the matrix directly surrounding new vascular cells . Other soluble proteases include MMPs-1, 2, 8, and 13 that are only activated in the extracellular matrix and degrade matrix components to enable neovessel growth . MMPs are inhibited by mediators known as tissue inhibitors of MMPs (TIMPs) that highly regulate the breakdown of matrix. Additionally, cysteine proteases known as cathepsins and serine proteases have been shown to control blood vessel formation. These complex interactions between cells and their matrix help facilitate neovascularization from the initial mobilization of EPCs to their ultimate fate as neovessels [109, 110]. Together, these studies indicate that remodeling enzymes such as MMP-9 and others play a crucial role in vasculogenesis at both the injury site and in the bone-marrow where quiescent EPCs reside.
The TGF superfamily consists of over 30 growth and differentiation factors that play vital roles in development and regulation of stem cell fate . During wound healing, specific TGF isoforms (1, 2, and 3) are secreted as a complex with latent precursors that are modified in the extracellular space. TGFβ is amongst the most well-studied signaling molecule in wound healing and is particularly linked to matrix and collagen production during wound healing. The ratio of expression of TGF1 and TGF3 is thought to regulate the ability of certain species and early gestation human fetuses to heal without scar .
In addition to its established role in fibrotic processes, TGF has been linked to neovascularization pathways through multiple receptor and intracellular signaling mechanisms. For example, TGF modulates vascular development by augmenting VEGF synthesis through Akt and ERK pathways . TGF has also been shown to activate the recruitment of VEGF-expressing hematopoietic effector cells, establishing a potent signaling network in the inflammatory wound environment that simultaneously stimulates neovascularization .
TGF pathways can also act independently of VEGF. In an embryonic stem cell vasculogenesis model, TGF was shown to stimulate neovessel growth via activin receptor-like kinase (ALK) receptors . Moreover, TGF can regulate non-endothelial cells during blood vessel maturation, specifically promoting vessel muscularization by stimulating MSC differentiation into pericytes . TGF can also activate the transdifferentiation of EPCs into myocytes (a process linked to pathologic intimal hyperplasia), highlighting the importance of tightly controlled cytokine pathways in vascular homeostasis .
Dysregulated TGF signaling has been linked to vascular pathology in humans. Mutations in the human endoglin gene, a TGF co-receptor, result in a vascular dysplasia known as hereditary hemorrhagic telangiectasia . EPCs from these patients exhibit aberrant ALK signaling and impaired vascular tubule formation in vitro, suggesting that TGF pathways are relevant to EPC function and vascular morphogenesis in humans. Collectively, these studies underscore the complex roles played by cytokines such as TGFβ in activating EPCs and mesenchymal precursors to produce functional neovasculature.
4. Challenges for Translation
A cascade of cytokines, growth factors, and other soluble mediators is released immediately following injury to orchestrate the repair of complex tissues . Numerous in vitro and preclinical studies have demonstrated that cytokine-based therapies can have a profound and multifaceted effect on neovascularization and chronic wound healing  (Figure 3). Although most of these therapies remain unproven in controlled clinical trials, several recombinant cytokines have been shown to have a positive impact on nonhealing wounds.
Recombinant human PDGF (becaplermin) is approved by the US Food and Drug Administration for the topical treatment of lower extremity diabetic neuropathic ulcers. Although several randomized controlled studies have validated its efficacy for nonhealing wounds, it remains expensive and not widely utilized [121, 122]. Granulocyte-CSF (G-CSF) is another cytokine that has demonstrated clinical benefit for diabetic patients with foot infections. G-CSF limits the duration of antibiotic treatment, hospital length of stay, and rate of amputation [123, 124]. Despite small case reports suggesting its effectiveness for chronic ulcers [125–129], larger clinical studies are needed to determine its ability to enhance wound healing.
Chemokine therapies are a promising strategy to promote neovascularization via modulation of the inflammatory response. Studies indicate that altered chemokine pathways may play a role in perpetuating the nonhealing nature of venous stasis ulcers . These wounds may also have ineffective angiogenic drives, suggesting that molecular strategies capable of augmenting blood vessel formation may prove clinically successful . For example, gene transfer of SDF-1 significantly enhanced EPC mobilization and vascularization in a hindlimb ischemia model, effects mediated through VEGF and nitric oxide synthase (NOS) . In a similar model, VEGF-transduced EPCs significantly improved wound vascularity compared to control EPCs , suggesting that EPC-targeted approaches may be a feasible option for clinical therapy.
In addition to gene-based therapies, biomaterial delivery of vasculogenic cytokines has been shown to improve vasculogenesis during wound healing [134, 135]. Matrix components and spatial patterning can precisely regulate vasculogenic programs and have the potential to promote a richly vascularized repair environment [136, 137]. As these biomaterial and molecular technologies continue to advance, combination cytokine-EPC impregnated scaffolds may become a clinical reality. Currently, wound therapies targeting vasculogenic pathways are largely in the preclinical stage but we believe these evolving strategies will continue to represent a promising approach to chronic wound healing.
Conflict of Interests
The authors have no financial conflict of interests to disclose.
- R. Kahn, R. M. Robertson, R. Smith, and D. Eddy, “The impact of prevention on reducing the burden of cardiovascular disease,” Circulation, vol. 118, no. 5, pp. 576–585, 2008.
- C. K. Sen, G. M. Gordillo, S. Roy et al., “Human skin wounds: a major and snowballing threat to public health and the economy,” Wound Repair and Regeneration, vol. 17, no. 6, pp. 763–771, 2009.
- S. C. Wu, V. R. Driver, J. S. Wrobel, and D. G. Armstrong, “Foot ulcers in the diabetic patient, prevention and treatment,” Vascular Health and Risk Management, vol. 3, no. 1, pp. 65–76, 2007.
- K. Stockl, A. Vanderplas, E. Tafesse, and E. Chang, “Costs of lower-extremity ulcers among patients with diabetes,” Diabetes Care, vol. 27, no. 9, pp. 2129–2134, 2004.
- J. B. Brown, K. L. Pedula, and A. W. Bakst, “The progressive cost of complications in type 2 diabetes mellitus,” Archives of Internal Medicine, vol. 159, no. 16, pp. 1873–1880, 1999.
- F. Carral, M. Aguilar, G. Olveira, A. Mangas, I. Doménech, and I. Torres, “Increased hospital expenditures in diabetic patients hospitalized for cardiovascular diseases,” Journal of Diabetes and its Complications, vol. 17, no. 6, pp. 331–336, 2003.
- G. D. Yancopoulos, S. Davis, N. W. Gale, J. S. Rudge, S. J. Wiegand, and J. Holash, “Vascular-specific growth factors and blood vessel formation,” Nature, vol. 407, no. 6801, pp. 242–248, 2000.
- S. M. Bauer, R. J. Bauer, and O. C. Velazquez, “Angiogenesis, vasculogenesis, and induction of healing in chronic wounds,” Vascular and Endovascular Surgery, vol. 39, no. 4, pp. 293–306, 2005.
- J. E. Ferguson, R. W. Kelley, and C. Patterson, “Mechanisms of endothelial differentiation in embryonic vasculogenesis,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 25, no. 11, pp. 2246–2254, 2005.
- N. Ferrara and R. S. Kerbel, “Angiogenesis as a therapeutic target,” Nature, vol. 438, no. 7070, pp. 967–974, 2005.
- Y. T. Ding, S. Kumar, and D. C. Yu, “The role of endothelial progenitor cells in tumour vasculogenesis,” Pathobiology, vol. 75, no. 5, pp. 265–273, 2008.
- M. R. Swift and B. M. Weinstein, “Arterial-venous specification during development,” Circulation Research, vol. 104, no. 5, pp. 576–588, 2009.
- J. Folkman, “Tumor angiogenesis: therapeutic implications,” The New England Journal of Medicine, vol. 285, no. 21, pp. 1182–1186, 1971.
- O. M. Tepper, J. M. Capla, R. D. Galiano et al., “Adult vasculogenesis occurs through in situ recruitment, proliferation, and tubulization of circulating bone marrow-derived cells,” Blood, vol. 105, no. 3, pp. 1068–1077, 2005.
- G. L. Semenza, “Vasculogenesis, angiogenesis, and arteriogenesis: mechanisms of blood vessel formation and remodeling,” Journal of Cellular Biochemistry, vol. 102, no. 4, pp. 840–847, 2007.
- T. Asahara, T. Murohara, A. Sullivan et al., “Isolation of putative progenitor endothelial cells for angiogenesis,” Science, vol. 275, no. 5302, pp. 964–967, 1997.
- P. Carmeliet, “Manipulating angiogenesis in medicine,” Journal of Internal Medicine, vol. 255, no. 5, pp. 538–561, 2004.
- T. Asahara, A. Kawamoto, and H. Masuda, “Concise review: circulating endothelial progenitor cells for vascular medicine,” Stem Cells, vol. 29, pp. 1650–1655, 2011.
- D. P. Sieveking, A. Buckle, D. S. Celermajer, and M. K. C. Ng, “Strikingly different angiogenic properties of endothelial progenitor cell subpopulations. Insights from a novel human angiogenesis assay,” Journal of the American College of Cardiology, vol. 51, no. 6, pp. 660–668, 2008.
- R. J. Medina, C. L. O'Neill, M. Sweeney et al., “Molecular analysis of endothelial progenitor cell (EPC) subtypes reveals two distinct cell populations with different identities,” BMC Medical Genomics, vol. 3, article no. 18, 2010.
- F. Timmermans, J. Plum, M. C. Yöder, D. A. Ingram, B. Vandekerckhove, and J. Case, “Endothelial progenitor cells: identity defined?” Journal of Cellular and Molecular Medicine, vol. 13, no. 1, pp. 87–102, 2009.
- K. Lamping, “Endothelial progenitor cells: sowing the seeds for vascular repair,” Circulation Research, vol. 100, no. 9, pp. 1243–1245, 2007.
- A. Zampetaki, J. P. Kirton, and Q. Xu, “Vascular repair by endothelial progenitor cells,” Cardiovascular Research, vol. 78, no. 3, pp. 413–421, 2008.
- C. Urbich, C. Heeschen, A. Aicher, et al., “Cathepsin L is required for endothelial progenitor cell-induced neovascularization,” Nature Medicine, vol. 11, pp. 206–213, 2005.
- X. W. Cheng, M. Kuzuya, K. Nakamura et al., “Mechanisms underlying the impairment of ischemia-induced neovascularization in matrix metalloproteinase 2-deficient mice,” Circulation Research, vol. 100, no. 6, pp. 904–913, 2007.
- S. Hamed, B. Brenner, A. Aharon, D. Daoud, and A. Roguin, “Nitric oxide and superoxide dismutase modulate endothelial progenitor cell function in type 2 diabetes mellitus,” Cardiovascular Diabetology, vol. 8, article no. 56, 2009.
- A. Iwakura, C. Luedemann, S. Shastry et al., “Estrogen-mediated, endothelial nitric oxide synthase-dependent mobilization of bone marrow-derived endothelial progenitor cells contributes to reendothelialization after arterial injury,” Circulation, vol. 108, no. 25, pp. 3115–3121, 2003.
- M. Crisan, S. Yap, L. Casteilla et al., “A perivascular origin for mesenchymal stem cells in multiple human organs,” Cell Stem Cell, vol. 3, no. 3, pp. 301–313, 2008.
- H. Gerhardt and C. Betsholtz, “Endothelial-pericyte interactions in angiogenesis,” Cell and Tissue Research, vol. 314, no. 1, pp. 15–23, 2003.
- F. Loffredo and R. T. Lee, “Therapeutic vasculogenesis: it takes two,” Circulation Research, vol. 103, no. 2, pp. 128–130, 2008.
- S. M. Bauer, R. J. Bauer, Z. J. Liu, H. Chen, L. Goldstein, and O. C. Velazquez, “Vascular endothelial growth factor-C promotes vasculogenesis, angiogenesis, and collagen constriction in three-dimensional collagen gels,” Journal of Vascular Surgery, vol. 41, no. 4, pp. 699–707, 2005.
- D. C. Rafii, B. Psaila, J. Butler, D. K. Jin, and D. Lyden, “Regulation of vasculogenesis by platelet-mediated recruitment of bone marrow-derived cells,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 28, no. 2, pp. 217–222, 2008.
- L. Coultas, K. Chawengsaksophak, and J. Rossant, “Endothelial cells and VEGF in vascular development,” Nature, vol. 438, no. 7070, pp. 937–945, 2005.
- R. Roskoski, “VEGF receptor protein-tyrosine kinases: structure and regulation,” Biochemical and Biophysical Research Communications, vol. 375, no. 3, pp. 287–291, 2008.
- J. A. Nagy, A. M. Dvorak, and H. F. Dvorak, “VEGF-A164/165 and PlGF: roles in angiogenesis and arteriogenesis,” Trends in Cardiovascular Medicine, vol. 13, no. 5, pp. 169–175, 2003.
- Y. Li, F. Zhang, N. Nagai et al., “VEGF-B inhibits apoptosis via VEGFR-1-mediated suppression of the expression of BH3-only protein genes in mice and rats,” Journal of Clinical Investigation, vol. 118, no. 3, pp. 913–923, 2008.
- T. Asahara, T. Takahashi, H. Masuda et al., “VEGF contributes to postnatal neovascularization by mobilizing bone marrow-derived endothelial progenitor cells,” The EMBO Journal, vol. 18, no. 14, pp. 3964–3972, 1999.
- B. Li, E. E. Sharpe, A. B. Maupin et al., “VEGF and PlGF promote adult vasculogenesis by enhancing EPC recruitment and vessel formation at the site of tumor neovascularization,” The FASEB Journal, vol. 20, no. 9, pp. 1495–1497, 2006.
- R. D. Galiano, O. M. Tepper, C. R. Pelo et al., “Topical vascular endothelial growth factor accelerates diabetic wound healing through increased angiogenesis and by mobilizing and recruiting bone marrow-derived cells,” American Journal of Pathology, vol. 164, no. 6, pp. 1935–1947, 2004.
- F. Caiado and S. Dias, “Endothelial progenitor cells and integrins: adhesive needs,” Fibrogenesis Tissue Repair, vol. 5, article 4, 2012.
- D. J. Ceradini and G. C. Gurtner, “Homing to hypoxia: HIF-1 as a mediator of progenitor cell recruitment to injured tissue,” Trends in Cardiovascular Medicine, vol. 15, no. 2, pp. 57–63, 2005.
- H. Thangarajah, D. Yao, E. I. Chang et al., “The molecular basis for impaired hypoxia-induced VEGF expression in diabetic tissues,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 32, pp. 13505–13510, 2009.
- M. R. Hoenig, C. Bianchi, and F. W. Sellke, “Hypoxia inducible factor -1α, endothelial progenitor cells, monocytes cardiovascular risk, wound healing, cobalt and hydralazine: a unifying hypothesis,” Current Drug Targets, vol. 9, no. 5, pp. 422–435, 2008.
- I. R. Botusan, V. G. Sunkari, O. Savu et al., “Stabilization of HIF-1α is critical to improve wound healing in diabetic mice,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 49, pp. 19426–19431, 2008.
- A. Peled, I. Petit, O. Kollet et al., “Dependence of human stem cell engraftment and repopulation of NOD/SCID mice on CXCR4,” Science, vol. 283, no. 5403, pp. 845–848, 1999.
- K. E. McGrath, A. D. Koniski, K. M. Maltby, J. K. McGann, and J. Palis, “Embryonic expression and function of the chemokine SDF-1 and its receptor, CXCR4,” Developmental Biology, vol. 213, no. 2, pp. 442–456, 1999.
- G. C. Gurtner, S. A. Loh, E. I. Chang et al., “SDF-1α expression during wound healing in the aged is HIF dependent,” Plastic and Reconstructive Surgery, vol. 123, pp. 65S–75S, 2009.
- D. J. Ceradini, A. R. Kulkarni, M. J. Callaghan et al., “Progenitor cell trafficking is regulated by hypoxic gradients through HIF-1 induction of SDF-1,” Nature Medicine, vol. 10, no. 8, pp. 858–864, 2004.
- J. I. Yamaguchi, K. F. Kusano, O. Masuo et al., “Stromal cell-derived factor-1 effects on ex vivo expanded endothelial progenitor cell recruitment for ischemic neovascularization,” Circulation, vol. 107, no. 9, pp. 1322–1328, 2003.
- K. I. Hiasa, M. Ishibashi, K. Ohtani et al., “Gene transfer of stromal cell-derived factor-1α enhances ischemic vasculogenesis and angiogenesis via vascular endothelial growth factor/endothelial nitric oxide synthase-related pathway: next-generation chemokine therapy for therapeutic neovascularization,” Circulation, vol. 109, no. 20, pp. 2454–2461, 2004.
- K. A. Gallagher, Z. J. Liu, M. Xiao et al., “Diabetic impairments in NO-mediated endothelial progenitor cell mobilization and homing are reversed by hyperoxia and SDF-1α,” Journal of Clinical Investigation, vol. 117, no. 5, pp. 1249–1259, 2007.
- D. Xing, L. Liu, G. P. Marti et al., “Hypoxia and hypoxia-inducible factor in the burn wound,” Wound Repair and Regeneration, vol. 19, no. 2, pp. 205–213, 2011.
- O. Z. Lerman, M. R. Greives, S. P. Singh et al., “Low-dose radiation augments vasculogenesis signaling through HIF-1-dependent and -independent SDF-1 induction,” Blood, vol. 116, no. 18, pp. 3669–3676, 2010.
- D. I. R. Holmes and I. Zachary, “The vascular endothelial growth factor (VEGF) family: angiogenic factors in health and disease,” Genome Biology, vol. 6, no. 2, article no. 209, 2005.
- S. Lange, J. Heger, G. Euler, M. Wartenberg, H. M. Piper, and H. Sauer, “Platelet-derived growth factor BB stimulates vasculogenesis of embryonic stem cell-derived endothelial cells by calcium-mediated generation of reactive oxygen species,” Cardiovascular Research, vol. 81, no. 1, pp. 159–168, 2009.
- M. Hellström, M. Kalén, P. Lindahl, A. Abramsson, and C. Betsholtz, “Role of PDGF-B and PDGFR-β in recruitment of vascular smooth muscle cells and pericytes during embryonic blood vessel formation in the mouse,” Development, vol. 126, no. 14, pp. 3047–3055, 1999.
- P. Lindahl, M. Hellström, M. Kalén et al., “Paracrine PDGF-B/PDGF-Rβ signaling controls mesangial cell development in kidney glomeruli,” Development, vol. 125, no. 17, pp. 3313–3322, 1998.
- S. G. Ball, C. A. Shuttleworth, and C. M. Kielty, “Mesenchymal stem cells and neovascularization: role of platelet-derived growth factor receptors,” Journal of Cellular and Molecular Medicine, vol. 11, no. 5, pp. 1012–1030, 2007.
- Q. Jin, G. Wei, Z. Lin et al., “Nanofibrous scaffolds incorporating PDGF-BB microspheres induce chemokine expression and tissue neogenesis in vivo,” PLoS ONE, vol. 3, no. 3, Article ID e1729, 2008.
- P. U. Magnusson, C. Looman, A. Åhgren, Y. Wu, L. Claesson-Welsh, and R. L. Heuchel, “Platelet-derived growth factor receptor-β constitutive activity promotes angiogenesis in vivo and in vitro,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 27, no. 10, pp. 2142–2149, 2007.
- J. Shen, M. D. Vil, M. Prewett et al., “Development of a fully human anti-PDGFRβ antibody that suppresses growth of human tumor xenografts and enhances antitumor activity of an anti-VEGFR2 antibody,” Neoplasia, vol. 11, no. 6, pp. 594–604, 2009.
- M. O. Murphy, J. Ghosh, P. Fulford et al., “Expression of growth factors and growth factor receptor in non-healing and healing ischaemic ulceration,” European Journal of Vascular and Endovascular Surgery, vol. 31, no. 5, pp. 516–522, 2006.
- R. K. Jain, “Molecular regulation of vessel maturation,” Nature Medicine, vol. 9, no. 6, pp. 685–693, 2003.
- M. C. Robson, D. P. Hill, P. D. Smith et al., “Sequential cytokine therapy for pressure ulcers: clinical and mechanistic response,” Annals of Surgery, vol. 231, no. 4, pp. 600–611, 2000.
- B. Thisse and C. Thisse, “Functions and regulations of fibroblast growth factor signaling during embryonic development,” Developmental Biology, vol. 287, no. 2, pp. 390–402, 2005.
- M. Murakami and M. Simons, “Fibroblast growth factor regulation of neovascularization,” Current Opinion in Hematology, vol. 15, no. 3, pp. 215–220, 2008.
- K. A. Detillieux, F. Sheikh, E. Kardami, and P. A. Cattini, “Biological activities of fibroblast growth factor-2 in the adult myocardium,” Cardiovascular Research, vol. 57, no. 1, pp. 8–19, 2003.
- M. L. Moya, S. Lucas, M. Francis-Sedlak et al., “Sustained delivery of FGF-1 increases vascular density in comparison to bolus administration,” Microvascular Research, vol. 78, no. 2, pp. 142–147, 2009.
- M. L. Moya, M. H. Cheng, J. J. Huang et al., “The effect of FGF-1 loaded alginate microbeads on neovascularization and adipogenesis in a vascular pedicle model of adipose tissue engineering,” Biomaterials, vol. 31, no. 10, pp. 2816–2826, 2010.
- H. Layman, M. Sacasa, A. E. Murphy, A. M. Murphy, S. M. Pham, and F. M. Andreopoulos, “Co-delivery of FGF-2 and G-CSF from gelatin-based hydrogels as angiogenic therapy in a murine critical limb ischemic model,” Acta Biomaterialia, vol. 5, no. 1, pp. 230–239, 2009.
- P. E. Burger, S. Coetzee, W. L. McKeehan et al., “Fibroblast growth factor receptor-1 is expressed by endothelial progenitor cells,” Blood, vol. 100, no. 10, pp. 3527–3535, 2002.
- M. Takahashi, N. Okubo, N. Chosa, et al., “Fibroblast growth factor-1-induced ERK1/2 signaling reciprocally regulates proliferation and smooth muscle cell differentiation of ligament-derived endothelial progenitor cell-like cells,” International Journal of Molecular Medicine, vol. 29, pp. 357–364, 2012.
- P. Dell'Era, M. Belleri, H. Stabile, M. L. Massardi, D. Ribatti, and M. Presta, “Paracrine and autocrine effects of fibroblast growth factor-4 in endothelial cells,” Oncogene, vol. 20, no. 21, pp. 2655–2663, 2001.
- A. Gualandris, M. Rusnati, M. Belleri et al., “Basic fibroblast growth factor overexpression in endothelial cells: an autocrine mechanism for angiogenesis and angioproliferative diseases,” Cell Growth and Differentiation, vol. 7, no. 2, pp. 147–160, 1996.
- J. Asai, H. Takenaka, M. Ii, et al., “Topical application of ex vivo expanded endothelial progenitor cells promotes vascularisation and wound healing in diabetic mice,” International Wound Journal, 2012.
- T. Takahashi, C. Kalka, H. Masuda et al., “Ischemia- and cytokine-induced mobilization of bone marrow-derived endothelial progenitor cells for neovascularization,” Nature Medicine, vol. 5, no. 4, pp. 434–438, 1999.
- A. Mann, K. Breuhahn, P. Schirmacher, and M. Blessing, “Keratinocyte-derived granulocyte-macrophage colony stimulating factor accelerates wound healing: Stimulation of keratinocyte proliferation, granulation tissue formation, and vascularization,” Journal of Investigative Dermatology, vol. 117, no. 6, pp. 1382–1390, 2001.
- X. Hu, H. Sun, C. Han, X. Wang, and W. Yu, “Topically applied rhGM-CSF for the wound healing: a systematic review,” Burns, vol. 37, no. 5, pp. 728–740, 2011.
- T. R. Hercus, D. Thomas, M. A. Guthridge et al., “The granulocyte-macrophage colony-stimulating factor receptor: linking its structure to cell signaling and its role in disease,” Blood, vol. 114, no. 7, pp. 1289–1298, 2009.
- T. D. Eubank, R. Roberts, M. Galloway, Y. Wang, D. E. Cohn, and C. B. Marsh, “GM-CSF induces expression of soluble VEGF receptor-1 from human monocytes and inhibits angiogenesis in mice,” Immunity, vol. 21, no. 6, pp. 831–842, 2004.
- K. Grote, H. Schuett, G. Salguero et al., “Toll-like receptor 2/6 stimulation promotes angiogenesis via GM-CSF as a potential strategy for immune defense and tissue regeneration,” Blood, vol. 115, no. 12, pp. 2543–2552, 2010.
- W. Zhao and X. H. Yang, “PI3K-dependent ERK is involved in GM-CSF-mediated activation of progenitor cells in a neovasculari sation model,” South African Journal of Science, vol. 106, no. 5-6, 2010.
- J. C. Kovacic, D. W. M. Muller, and R. M. Graham, “Actions and therapeutic potential of G-CSF and GM-CSF in cardiovascular disease,” Journal of Molecular and Cellular Cardiology, vol. 42, no. 1, pp. 19–33, 2007.
- R. Shen, Y. Ye, L. Chen, Q. Yan, S. H. Barsky, and J. X. Gao, “Precancerous stem cells can serve as tumor vasculogenic progenitors,” PLoS ONE, vol. 3, no. 2, Article ID e1652, 2008.
- R. Estrada, Q. Zeng, H. Lu et al., “Up-regulating sphingosine 1-phosphate receptor-2 signaling impairs chemotactic, wound-healing, and morphogenetic responses in senescent endothelial cells,” The Journal of Biological Chemistry, vol. 283, no. 44, pp. 30363–30375, 2008.
- R. Vogler, B. Sauer, D. S. Kim, M. Schäfer-Korting, and B. Kleuser, “Sphingosine-1-phosphate and its potentially paradoxical effects on critical parameters of cutaneous wound healing,” Journal of Investigative Dermatology, vol. 120, no. 4, pp. 693–700, 2003.
- T. Kawanabe, T. Kawakami, Y. Yatomi, S. Shimada, and Y. Soma, “Sphingosine 1-phosphate accelerates wound healing in diabetic mice,” Journal of Dermatological Science, vol. 48, no. 1, pp. 53–60, 2007.
- Y. Hisano, N. Kobayashi, A. Yamaguchi, and T. Nishi, “Mouse SPNS2 functions as a sphingosine-1-phosphate transporter in vascular endothelial cells,” PloS One, vol. 7, Article ID e38941, 2012.
- C. Givens and E. Tzima, “S1P1 bridges mechanotransduction and angiogenesis during vascular development,” Developmental Cell, vol. 23, pp. 451–452, 2012.
- C. Tobia, P. Chiodelli, S. Nicoli, et al., “Sphingosine-1-phosphate receptor-1 controls venous endothelial barrier integrity in zebrafish,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 32, pp. e104–e116, 2012.
- K. M. Argraves, B. A. Wilkerson, W. S. Argraves, P. A. Fleming, L. M. Obeid, and C. J. Drake, “Sphingosine-1-phosphate signaling promotes critical migratory events in vasculogenesis,” The Journal of Biological Chemistry, vol. 279, no. 48, pp. 50580–50590, 2004.
- Y. Liu, R. Wada, T. Yamashita et al., “Edg-1, the G protein-coupled receptor for sphingosine-1-phosphate, is essential for vascular maturation,” Journal of Clinical Investigation, vol. 106, no. 8, pp. 951–961, 2000.
- H. M. Rosenfeldt, J. P. Hobson, S. Milstien, and S. Spiegel, “The sphingosine-1-phosphate receptor EDG-1 is essential for platelet-derived growth factor-induced cell motility,” Biochemical Society Transactions, vol. 29, no. 6, pp. 836–839, 2001.
- J. A. Manuel and B. Gawronska-Kozak, “Matrix metalloproteinase 9 (MMP-9) is upregulated during scarless wound healing in athymic nude mice,” Matrix Biology, vol. 25, no. 8, pp. 505–514, 2006.
- Y. Liu, D. Min, T. Bolton et al., “Increased matrix metalloproteinase-9 predicts poor wound healing in diabetic foot ulcers,” Diabetes Care, vol. 32, no. 1, pp. 117–119, 2009.
- T. R. Kyriakides, D. Wulsin, E. A. Skokos et al., “Mice that lack matrix metalloproteinase-9 display delayed wound healing associated with delayed reepithelization and disordered collagen fibrillogenesis,” Matrix Biology, vol. 28, no. 2, pp. 65–73, 2009.
- V. Masson, L. Rodriguez De La Ballina, C. Munaut et al., “Contribution of host MMP-2 and MMP-9 to promote tumor vascularization and invasion of malignant keratinocytes,” The FASEB Journal, vol. 19, no. 2, pp. 234–236, 2005.
- D. Stenzel, C. A. Franco, S. Estrach, et al., “Endothelial basement membrane limits tip cell formation by inducing Dll4/Notch signalling in vivo,” EMBO Report, vol. 12, pp. 1135–1143, 2011.
- A. N. Stratman, K. M. Malotte, R. D. Mahan, M. J. Davis, and G. E. Davis, “Pericyte recruitment during vasculogenic tube assembly stimulates endothelial basement membrane matrix formation,” Blood, vol. 114, no. 24, pp. 5091–5101, 2009.
- G. E. Davis and D. R. Senger, “Endothelial extracellular matrix: biosynthesis, remodeling, and functions during vascular morphogenesis and neovessel stabilization,” Circulation Research, vol. 97, no. 11, pp. 1093–1107, 2005.
- B. Heissig, K. Hattori, M. Friedrich, S. Rafii, and Z. Werb, “Angiogenesis: vascular remodeling of the extracellular matrix involves metalloproteinases,” Current Opinion in Hematology, vol. 10, no. 2, pp. 136–141, 2003.
- B. Heissig, K. Hattori, S. Dias et al., “Recruitment of stem and progenitor cells from the bone marrow niche requires MMP-9 mediated release of Kit-ligand,” Cell, vol. 109, no. 5, pp. 625–637, 2002.
- G. Bergers, R. Brekken, G. McMahon et al., “Matrix metalloproteinase-9 triggers the angiogenic switch during carcinogenesis,” Nature Cell Biology, vol. 2, no. 10, pp. 737–744, 2000.
- Q. Yu and I. Stamenkovic, “Cell surface-localized matrix metalloproteinase-9 proteolytically activates TGF-β and promotes tumor invasion and angiogenesis,” Genes and Development, vol. 14, no. 2, pp. 163–176, 2000.
- G. O. Ahn and J. M. Brown, “Matrix metalloproteinase-9 is required for tumor vasculogenesis but not for angiogenesis: role of bone marrow-derived myelomonocytic cells,” Cancer Cell, vol. 13, no. 3, pp. 193–205, 2008.
- P. H. Huang, Y. H. Chen, C. H. Wang et al., “Matrix metalloproteinase-9 is essential for ischemia-induced neovascularization by modulating bone marrow-derived endothelial progenitor cells,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 29, no. 8, pp. 1179–1184, 2009.
- P. E. Szmitko, P. W. M. Fedak, R. D. Weisel, D. J. Stewart, M. J. B. Kutryk, and S. Verma, “Endothelial progenitor cells: new hope for a broken heart,” Circulation, vol. 107, no. 24, pp. 3093–3100, 2003.
- V. W. M. Van Hinsbergh, M. A. Engelse, and P. H. A. Quax, “Pericellular proteases in angiogenesis and vasculogenesis,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 26, no. 4, pp. 716–728, 2006.
- V. W. M. Van Hinsbergh and P. Koolwijk, “Endothelial sprouting and angiogenesis: matrix metalloproteinases in the lead,” Cardiovascular Research, vol. 78, no. 2, pp. 203–212, 2008.
- J. E. Rundhaug, “Matrix metalloproteinases and angiogenesis,” Journal of Cellular and Molecular Medicine, vol. 9, no. 2, pp. 267–285, 2005.
- K. Kitisin, T. Saha, T. Blake et al., “Tgf-Beta signaling in development,” Science's STKE, vol. 2007, no. 399, p. cm1, 2007.
- B. J. Larson, M. T. Longaker, and H. P. Lorenz, “Scarless fetal wound healing: a basic science review,” Plastic and Reconstructive Surgery, vol. 126, no. 4, pp. 1172–1180, 2010.
- P. ten Dijke and H. M. Arthur, “Extracellular control of TGF-beta signalling in vascular development and disease,” Nature Reviews Molecular Cell Biology, vol. 8, pp. 857–869, 2007.
- S. Fang, N. Pentinmikko, M. Ilmonen, and P. Salven, “Dual action of TGF-beta induces vascular growth in vivo through recruitment of angiogenic VEGF-producing hematopoietic effector cells,” Angiogenesis, vol. 15, pp. 511–519, 2012.
- C. Mallet, D. Vittet, J. J. Feige, and S. Bailly, “TGFβ1 induces vasculogenesis and inhibits angiogenic sprouting in an embryonic stem cell differentiation model: respective contribution of ALK1 and ALK5,” Stem Cells, vol. 24, no. 11, pp. 2420–2427, 2006.
- X.-J. Wang, Z. Dong, X.-H. Zhong, et al., “Transforming growth factor-beta1 enhanced vascular endothelial growth factor synthesis in mesenchymal stem cells,” Biochemical and Biophysical Research Communications, vol. 365, pp. 548–554, 2008.
- H. Imamura, T. Ohta, K. Tsunetoshi et al., “Transdifferentiation of bone marrow-derived endothelial progenitor cells into the smooth muscle cell lineage mediated by tansforming growth factor-β1,” Atherosclerosis, vol. 211, no. 1, pp. 114–121, 2010.
- L. A. Fernandez, F. Sanz-Rodriguez, F. J. Blanco, C. Bernabéu, and L. M. Botella, “Hereditary hemorrhagic telangiectasia, a vascular dysplasia affecting the TGF-β signaling pathway,” Clinical Medicine and Research, vol. 4, no. 1, pp. 66–78, 2006.
- S. Werner and R. Grose, “Regulation of wound healing by growth factors and cytokines,” Physiological Reviews, vol. 83, no. 3, pp. 835–870, 2003.
- S. Barrientos, O. Stojadinovic, M. S. Golinko, H. Brem, and M. Tomic-Canic, “Growth factors and cytokines in wound healing,” Wound Repair and Regeneration, vol. 16, no. 5, pp. 585–601, 2008.
- R. C. Fang and R. D. Galiano, “A review of becaplermin gel in the treatment of diabetic neuropathic foot ulcers,” Biologics, vol. 2, pp. 1–12, 2008.
- N. Papanas and E. Maltezos, “Becaplermin gel in the treatment of diabetic neuropathic foot ulcers,” Clinical Interventions in Aging, vol. 3, no. 2, pp. 233–240, 2008.
- A. Gough, M. Clapperton, N. Rolando, A. V. M. Foster, J. Philpott-Howard, and M. E. Edmonds, “Randomised placebo-controlled trial of granulocyte-colony stimulating factor in diabetic foot infection,” The Lancet, vol. 350, no. 9081, pp. 855–859, 1997.
- F. De Lalla, G. Pellizzer, M. Strazzabosco et al., “Randomized prospective controlled trial of recombinant granulocyte colony-stimulating factor as adjunctive therapy for limb-threatening diabetic foot infection,” Antimicrobial Agents and Chemotherapy, vol. 45, no. 4, pp. 1094–1098, 2001.
- R. Marques Da Costa, C. Aniceto, F. Miguel Jesus, and M. Mendes, “Quick healing of leg ulcers after molgramostim,” The Lancet, vol. 344, no. 8920, pp. 481–482, 1994.
- Z. Pojda and J. Struzyna, “Treatment of non-healing ulcers with rhGM-CSF and skin grafts,” The Lancet, vol. 343, no. 8905, p. 1100, 1994.
- M. Raderer, G. Kornek, M. Hejna, K. Koperna, W. Scheithauer, and W. Base, “Topical granulocyte-macrophage colony-stimulating factor in patients with cancer and impaired wound healing,” Journal of the National Cancer Institute, vol. 89, no. 3, p. 263, 1997.
- F. Arnold, J. O'Brien, and G. Cherry, “Granulocyte monocyte-colony stimulating factor as an agent for wound healing. A study evaluating the use of local injections of a genetically engineered growth factor in the management of wounds with a poor healing prognosis,” Journal of Wound Care, vol. 4, no. 9, pp. 400–402, 1995.
- R. Marques Da Costa, F. M. Jesus, C. Aniceto, and M. Mendes, “Double-blind randomized placebo-controlled trial of the use of granulocyte-macrophage colony-stimulating factor in chronic leg ulcers,” American Journal of Surgery, vol. 173, no. 3, pp. 165–168, 1997.
- D. P. Fivenson, D. T. Faria, B. J. Nickoloff et al., “Chemokine and inflammatory cytokine changes during chronic wound healing,” Wound Repair and Regeneration, vol. 5, no. 4, pp. 310–322, 1997.
- S. L. Drinkwater, K. G. Burnand, R. Ding, and A. Smith, “Increased but ineffectual angiogenic drive in nonhealing venous leg ulcers,” Journal of Vascular Surgery, vol. 38, no. 5, pp. 1106–1112, 2003.
- K. I. Hiasa, M. Ishibashi, K. Ohtani et al., “Gene transfer of stromal cell-derived factor-1α enhances ischemic vasculogenesis and angiogenesis via vascular endothelial growth factor/endothelial nitric oxide synthase-related pathway: next-generation chemokine therapy for therapeutic neovascularization,” Circulation, vol. 109, no. 20, pp. 2454–2461, 2004.
- H. Iwaguro, J. I. Yamaguchi, C. Kalka et al., “Endothelial progenitor cell vascular endothelial growth factor gene transfer for vascular regeneration,” Circulation, vol. 105, no. 6, pp. 732–738, 2002.
- S. Y. Rabbany, J. Pastore, M. Yamamoto et al., “Continuous delivery of stromal cell-derived factor-1 from alginate scaffolds accelerates wound healing,” Cell Transplantation, vol. 19, no. 4, pp. 399–408, 2010.
- P. W. Henderson, S. P. Singh, D. D. Krijgh et al., “Stromal-derived factor-1 delivered via hydrogel drug-delivery vehicle accelerates wound healing in vivo,” Wound Repair and Regeneration, vol. 19, no. 3, pp. 420–425, 2011.
- D. Hanjaya-Putra, K. T. Wong, K. Hirotsu, S. Khetan, J. A. Burdick, and S. Gerecht, “Spatial control of cell-mediated degradation to regulate vasculogenesis and angiogenesis in hyaluronan hydrogels,” Biomaterials, vol. 33, pp. 6123–6131, 2012.
- D. Hanjaya-Putra, V. Bose, Y. I. Shen et al., “Controlled activation of morphogenesis to generate a functional human microvasculature in a synthetic matrix,” Blood, vol. 118, no. 3, pp. 804–815, 2011.