Stem Cells
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

First published online September 6, 2007
Stem Cells Vol. 25 No. 12 December 2007, pp. 3101 -3110
doi:10.1634/stemcells.2006-0795; www.StemCells.com
© 2007 AlphaMed Press

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
2006-0795v1
2006-0795v2
25/12/3101    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Reprints/Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Grenier, G.
Right arrow Articles by Rudnicki, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Grenier, G.
Right arrow Articles by Rudnicki, M. A.

TISSUE-SPECIFIC STEM CELLS

Resident Endothelial Precursors in Muscle, Adipose, and Dermis Contribute to Postnatal Vasculogenesis

Guillaume Greniera, Anthony Scimèa, Fabien Le Granda, Atsushi Asakuraa, Carolina Perez-Iratxetaa, Miguel A. Andrade-Navarroa, Patricia A. Laboskyb, Michael A. Rudnickia

aSprott Centre for Stem Cell Research, Ottawa Health Research Institute, Ottawa, Ontario, Canada;
bVanderbilt Center for Stem Cell Biology, Vanderbilt University, Nashville, Tennessee, USA

Key Words. Vasculogenesis • Endothelial precursors • Sca1 • Skeletal muscle

Correspondence: Michael A. Rudnicki, Ph.D., Sprott Centre for Stem Cell Research, Ottawa Health Research Institute, 501 Smyth Road, Ottawa, Ontario K1H 8L6, Canada. Telephone: 613-739-6737; Fax: 613-737-8803; e-mail: mrudnicki{at}ohri.ca

Received on December 20, 2006; accepted for publication on August 14, 2007.

First published online in STEM CELLS EXPRESS  September 6, 2007.

   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
A novel population of tissue-resident endothelial precursors (TEPs) was isolated from small blood vessels in dermal, adipose, and skeletal muscle of mouse based on their ability to be grown as spheres. Cellular and molecular analyses of these cells revealed that they were highly related regardless of the tissue of origin and distinct from embryonic neural stem cells. Notably, TEPs did not express hematopoietic markers, but they expressed numerous characteristics of angiogenic precursors and their differentiated progeny, such as CD34, Flk-1, Tie-1, CD31, and vascular endothelial cadherin (VE-cadherin). TEPs readily differentiated into endothelial cells in newly formed vascular networks following transplantation into regenerating skeletal muscle. Taken together, these experiments suggest that TEPs represent a novel class of endothelial precursors that are closely associated with small blood vessels in muscle, adipose, and dermal tissue. This finding is of particular interest since it could bring new insight in cancer angiogenesis and collateral blood vessels developed following ischemia.

Disclosure of potential conflicts of interest is found at the end of this article.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
The development of capillaries occurs during embryogenesis via the proliferation, specification, and differentiation of hemangioblast-derived cells into endothelial cells (EC) [1]. In healthy adult individuals, the low basal turnover of the endothelium (angiogenesis) is thought to occur by proliferation and migration of pre-existing mature EC [2–4]. However, during acute neovascularization (vasculogenesis), their capacity to reconstruct endothelium is limited, as mature EC possess a low proliferative potential [5].

Recent studies have revealed the presence of a bone marrow (BM)-derived endothelial progenitor cell (EPC) in systemic circulation [6]. Although the contribution of these EPC to angiogenesis is negligible, models of vasculogenesis such as limb ischemia or severe burns have demonstrated that circulating EPC can be actively recruited, despite the fact that they are found in relatively low numbers in peripheral blood [7–9]. More recently, monocyte/macrophage cells have been shown to acquire an endothelial phenotype when exposed to angiogenic conditions [10–13]. Interestingly, monocytes that express both CD45, a hematopoietic marker, and VE-cadherin, an endothelial marker, have been identified in neovascularized tumors [14]. However, EPC but not monocytic-derived endothelial progenitors are more efficient at angiogenesis in a limb ischemia mouse model [15]. It has remained unclear as to whether endothelial progenitors resident in tissues contribute in a significant manner to angiogenesis and vasculogenesis during regeneration following damage.

Several laboratories have described EC progenitors potentially resident in tissue. Isolation of CD34+CD31– or CD34–CD31– cells from the stromal vascular fraction of adipose tissue revealed that both populations differentiated in vitro into EC, promoted angiogenesis, and participated in the revascularization of ischemic hind limbs of nude mice [16, 17]. Other tissue-resident EC progenitors have been found in the heart by isolation of c-kit+ stem cells and in neural tissue using stem clones derived from coculture with EC [18, 19]. Side-population cells isolated from muscle tissue (mSP) on the basis of Hoechst dye exclusion when transplanted into regenerating muscle have been documented to engraft into endothelium during vascular regeneration [20].

mSP can reconstitute the blood system of irradiated mice following bone-marrow transplantation as well as participate in the regeneration of skeletal muscle [21, 22]. Most mSP possess the cell surface marker Sca1, which has previously been shown to be present on stem cell populations that give rise to muscle and endothelial cells [23, 24]. However, these studies did not investigate the physiological contribution of mSP toward either angiogenesis or vasculogenesis.

We set out to characterize a population of cells isolated from of mouse dermis, adipose, and muscle tissue that are cultured as spheres. We first identified the source of the cells within these tissues, investigated their developmental origin, and characterized their gene expression profile. Second, we characterized their differentiation potential both in vitro and in vivo. Third, we assessed their biological contribution during wound healing. Our experiments indicate that these cells represent a novel class of tissue-resident endothelial precursors (TEPs) closely associated with small blood vessels that actively participate in angiogenesis and vasculogenesis during tissue regeneration. The ready accessibility of TEPs from a variety of tissues, together with the potential to expand the precursor cell in vitro, raises the possibility of exploiting them for therapy and for the development of engineered tissues.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
This investigation was conducted in accordance with the rules and regulations of the Animal Care Committee of the University of Ottawa.

Cell Isolation and Culture
Dermal tissue from thoracic and abdominal skin was separated from epidermis by overnight incubation in 500 µg/ml thermolysin (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com) and digested with 0.1% trypsin (Invitrogen, Carlsbad, CA, http://www.invitrogen.com)/0.01% EDTA (Sigma-Aldrich) for 60 minutes at 37°C. Adipose and muscle tissues were digested with 1% collagenase I (Sigma-Aldrich) for 45 minutes at 37°C and 1% collagenase B (Roche Applied Science, Penzberg, Germany, http://www.roche-applied-science.com)/24 U/ml dispase (Grade II; Roche Diagnostics) for 12 minutes at 37°C, respectively. The tissue digests were washed with Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS) and penicillin/streptomycin, poured through a 100-µm cell strainer (BD Falcon, Mississauga, Canada, http://www.bdbiosciences.com). After centrifugation, cells were washed and resuspended in sphere-growing medium (SGM) (i.e., murine Neurocult basal medium [Stem Cell Technologies, Vancouver, BC, Canada, http://www.stemcell.com] supplemented with 1% B-27 [Invitrogen], 20 ng/ml epidermal growth factor [Stem Cell Technologies], 40 ng/ml basic fibroblast growth factor [bFGF; eBioshop, San Diego, CA, http://www.ebioscience.com], 20 ng/ml leukemia inhibitory factor [Chemicon, Temecula, CA, http://www.chemicon.com], and antibiotics). Resuspended cells were cultured into 75-cm2 straight-neck tissue culture flasks (BD Falcon) at 37°C and 5% CO2.

After 4 days, free-floating spheres, or TEPs, were transferred into a 15-ml tube and incubated for 2 minutes to let cells settle down. The supernatant, which contained single cells and debris, was discarded, and the pellet was mechanically dissociated with a fire-polished Pasteur pipette. The dissociated pellet was resuspended in fresh medium and reseeded into a 75-cm2 tissue culture flask. Cells were passaged every 6 days and used between passages 2 and 4.

TEPs were also purified using flow-activated cell sorting (FACS). For FACS, cells were incubated on ice for 30 minutes with 1 µg of antibody per 1 x 106 cells: Phycoerythrin (PE)-CD31 (BD Pharmingen, San Diego, http://www.bdbiosciences.com/pharmingen), PE-anti-Sca1 (BD Pharmingen), APC-Sca1 (Cedarlane), and biotinylated lineage (Lin) antibodies (BD Pharmingen) followed by fluorescein isothiocyanate (FITC)-conjugated-streptavidin secondary antibody. Analysis and/or sorting were performed on a MoFlo cytometer (DakoCytomation, Glostrup, Denmark, http://www.dakocytomation.com). The side population (SP) cell fraction was isolated as described elsewhere [22].

To isolate TEPs from small blood vessels, tissues were minced and then incubated for 15 minutes at 37°C with 1% collagenase I. After incubation, tissues were further digested by 500 µg/ml thermolysin in HEPES supplemented with 1 mM CaCl2 for 3 hours. Fragments of vessels were isolated with a coated Pasteur pipette, washed by serial transfer into Petri dishes containing fresh SGM medium, and finally sedimented for 2 minutes. Small blood vessels were resuspended with collagenase/dispase for 8 minutes at 37°C, and single-cell suspensions were obtained by trituration.

For neuronal differentiation, cells were plated in SGM at high density on two-well culture slides coated with poly-D-lysine/laminin (Biocoat; Becton, Dickinson and Company, Franklin Lakes, NJ, http://www.bdbiosciences.com). After 24 hours, SGM was replaced by neuronal differentiation medium (Stem Cell Technologies) for 4 days. For adipocyte differentiation, 90% confluent cells were resuspended in DMEM supplemented with 10% FBS, 1 µM dexamethasone (Sigma-Aldrich), 5 µg/ml insulin (Sigma-Aldrich) and 0.5 mM 3-isobutyl-1-methylxanthine (Sigma-Aldrich). After 48 hours, medium was replaced by DMEM supplemented with 10% FBS and 5 µg/ml insulin for 7 days. For smooth muscle cell differentiation, cells were cultured in DMEM with 2% horse serum for 5 days. For skeletal muscle cell differentiation, cells were cultured in Ham's F-10 medium containing 20% FBS and 2.5 ng/ml bFGF on collagen coated tissue culture flasks. At 80% confluence, medium was replaced with DMEM containing 5% horse serum. For endothelial cell differentiation, cells were plated on either collagen-coated or noncoated tissue culture flasks with endothelial growth medium 2 (Cambrex, Walkersville, MD, http://www.cambrex.com). To test the ability of isolated cells to uptake low-density lipoprotein (LDL), we incubated them with fluorescent 1,1'-dioctadecyl-3,3,3',3'-tetramethyl-indocarbocyanine acetylated LDL (Biomedical Technologies Incorporated (BTI), Stoughton, MA, www.btiinc.com) as described by the manufacturer. To assess capillary-like formation, TEPs were plated on Matrigel for 3 days and stained with alkaline phosphatase (ALKP) according to the manufacturer's instructions (Sigma-Aldrich). 5-Bromo-4-chloro-3-indolyl-β-D-galactoside (X-Gal) staining performed on Flt-1/lacZ-derived TEPs was described previously [25].

RNA, Polymerase Chain Reaction, and Microarray Analysis
RNA from three individual experiments was isolated using the RNeasy RNA isolation kit (Qiagen, Hilden, Germany, http://www.qiagen.com). For semiquantitative reverse transcription-polymerase chain reaction (RT-PCR), cDNA was first synthesized using 500 ng of RNA with the GeneAmp RNA PCR Core kit (Roche Diagnostics) according to the manufacturer's instructions. PCRs for different cDNA concentrations were 35 cycles at an annealing temperature of 55°C using previously described primers [26–30] or designed using web-available software Xpression Primer3 (Primer 3, Cambridge, MA, http://frodo.wi.mit.edu) (Sca1 forward, TGGATTCTCAAACAAGGAAAGTAAAGA; Sca1 reverse, ACCCAGGATCTCCATACTTTCAATA; CD45 forward, CCACCAGGGACTGACAAGTT, CD45 reverse, TAGGCTTAGGCGTTTCTGGA; VE-cadherin forward, CGGTCAAGTATGGGCAGTTT, VE-cadherin reverse, CAACTGCTCGTGAATCTCCA).

Biological triplicates of 10T1/2, embryo-derived neural stem cells (eNSC), and TEPs from adult dermal, muscle, and adipose tissues were analyzed by microarray as described in the Affymetrix GeneChip protocol (Affymetrix, Santa Clara, CA, http://www.affymetrix.com). Probes were hybridized to an identical lot of Affymetrix MOE430A and MOE430B GeneChip arrays, and the signal was determined using the MAS 5.0 absolute analysis algorithm, as the average fluorescence intensity among the intensities obtained from the probe set. Gene expression was analyzed using cluster V3.0 and Maple Tree 0,2,3.3 (http://rana.lbl.gov/EisenSoftware.htm).

Immunocytochemistry
Cells were fixed with 4% paraformaldehyde for 10 minutes and permeabilized with 0.3% Triton X-100 in phosphate-buffered saline. Cells were then incubated with primary antibodies: anti-desmin (Dako, Glostrup, Denmark, http://www.dako.com), anti-myoD (BD Pharmingen), anti-nestin (Stem Cell Technologies), anti-glial fibrillary acidic protein (anti-GFAP; Stem Cell Technologies), anti-β-III-tubulin (Stem Cell Technologies), anti-CD31 (BD Pharmingen), and anti-VE-cadherin (BD Pharmingen) overnight at 4°C. After incubation, cells were washed, and secondary antibody FITC-conjugated goat anti-mouse (Chemicon) or PE-conjugated goat anti-rabbit (Chemicon) was added for 30 minutes at 37°C. For CD31-labeled cells, biotin-conjugated anti-rat antibody was added at room temperature for 45 minutes, followed by incubation with avidin-Alexa 546-conjugated antibody (Molecular Probes Inc., Eugene, OR, http://probes.invitrogen.com) for 30 minutes at 37°C.

Biological Contribution of TEPs to Muscle Regeneration
Injury was induced by injecting 25 µl of 10 µM cardiotoxin (CTX; Latoxan, Valence, France, http://www.latoxan.com) into the host tibialis anterior (TA) muscle of Balb/c mice. Four days postinjury, 100,000 green fluorescent protein (GFP)+Sca1+CD31– donor cells from Tg(GFPU)5Nagy/J mice were injected into the host TA. Ten days post-transplantation, the muscle was digested and GFP+ cells were analyzed for the expression of CD31. In addition, Sca1+CD31– and eNSC-derived from Flt1/LacZ mice were also transplanted into the TA muscle of mice 4 days following CTX insult and analyzed after 10 days by whole mount staining with X-Gal to visualize the expression of β-galactosidase. To assess the biological contribution of endogenous TEPs during wound healing, TA muscles from CTX-injected and control mice were harvested 4 days post-treatment, weighed, and digested, and cells were counted. Counted cells were Sca1+CD31–Lin–-sorted, and 5,000 cells per cm2 were cultured in SGM. Ten days after growth in SGM, cells that formed spheres representing TEPs were counted, plated on Matrigel, and stained for ALKP.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Adult Sphere-Forming Cells Are Derived from Small Blood Vessels
As neural progenitors can be readily propagated in cell culture in SGM, we wanted to characterize other putative adult progenitor cells similarly propagated from different tissues [31, 32]. We first determined that spheres from dermis, adipose, and skeletal muscle tissue suspended in SGM could be isolated and propagated for few passages (data not shown).

We then assessed the localization of the cells that form spheres within the tissue of origin to find out whether these cells represent the same cell types, regardless of tissue derivation. Observational evidence suggested that they might be associated with small blood vessels by the vivid red appearance of pelleted cells before SGM culture after adipose, dermal, or muscle tissue digestion (Fig. 1A). To test this hypothesis, small blood vessels were collected manually after sequential digestion using collagenase and thermolysin (Fig. 1A). The collected structures were further broken down to obtain single-cell suspensions that were tested for their ability to give rise to spheres following SGM growth. We observed that the cells that formed spheres grew only from the small blood vessel-derived cell preparations of dermis, muscle, and adipose tissues but never from any of the non-blood vessel-derived tissue fractions (Fig. 1A).


Figure 1
View larger version (39K):
[in this window]
[in a new window]

  		Figure 1. Spheres are derived from mesoderm and are associated with the Sca1+CD31–Lin– cell fraction of blood vessels. (A): After tissue digestion, spheres were grown from a cell pellet potentially enriched with blood vessels. Only single cells originating from blood vessel preparations gave rise to spheres. (B): Reverse transcription-polymerase chain reaction for Sca1, CD45 VE-cadherin, and β-actin of muscle tissue; its freshly sorted Sca1+CD31–Lin– cell population; and differentiated Sca1+CD31–Lin– cells. Note that there was no cross-contamination by hematopoietic and mature endothelial cells during sphere isolation. (C): Immunostaining for VE-cadherin on undifferentiated and differentiated spheres. Note that VE-cadherin expression was present only in differentiated cells. (D): Schematic diagram displaying scheme for sphere isolation. Only total cells and the Sca1+CD31–Lin– fraction from blood vessels gave rise to spheres that differentiated into endothelial cells. (E): β-Galactosidase activity staining of spheres isolated from the muscle of Wnt-1-Cre x R26R and Tie-2-Cre x R26R mice. β-Galactosidase activity was detected only in spheres derived from Tie-2-cre x R26R mice, indicating a mesodermal origin. Abbreviation: rt, reverse transcriptase.

 
We next examined potential cell surface markers based on each sphere's blood vessel association that could be used in isolation and transplantation studies. As stem cell antigen-1 (Sca1) is predominately expressed on cells surrounding blood vessels we first sorted Sca1+ and Sca1– fractions of dermal, adipose and skeletal muscle tissues [22, 33]. After 4 days in culture, spheres appeared only in the Sca1+ population, but not in the Sca1– population (Table 1). Notably, it was also possible to grow spheres from the SP+ cell fraction that was enriched with Sca1+ cells (Table 1).


View this table:
[in this window]
[in a new window]

  		Table 1. Spheres originate from the Sca1+CD31–Lin– and SP+ cellular fractions of dermal, adipose, and skeletal muscle tissue

 
The cell surface marker characterization was further defined by fractionating the Sca1+ population with CD31 (or platelet-endothelial cell adhesion molecule), a mature endothelial marker. Spheres were isolated only in the CD31– fraction containing immature endothelial cells, regardless of tissue derivation (Table 1). This Sca1+CD31– cell-sorted population that formed spheres was also delineated by isolation of a segment of cells against a cocktail of antibodies recognizing the leukocyte lineage cells (Lin). Spheres arose only from the Sca1+CD31–Lin– cells and not the Sca1+CD31–Lin+ cells, suggesting that the spheres are not derived from hematopoietic precursors (Table 1). Moreover, we found that spheres could never be cultured from the mononuclear fraction of blood from which EPC are derived or from endothelial cells grown in vitro (Table 1).

To ensure that isolated spheres were not derived from hematopoietic or mature endothelial cells, we performed RT-PCR analyses for the expression of CD45, a hematopoietic cell marker, and CD144 (VE-cadherin), a marker for mature endothelial cells, on digested muscle tissue; the freshly sorted Sca1+CD31–Lin– cellular fraction, and the Sca1+CD31–Lin– cells cultured in endothelial cell differentiation medium (Fig. 1B). Our results demonstrated that CD45, a hematopoietic cell marker, was expressed in digested tissue but not in the sorted cells before and after differentiation, thereby excluding the possible hematopoietic cell cross-contamination. In addition, VE-cadherin, a mature endothelial cell marker, was expressed as expected in digested tissue but not in the Sca1+CD31–Lin– cell population. Interestingly, VE-cadherin was re-expressed in the sorted cells that were cultured for 4 days in endothelial differentiation media (Fig. 1B). We confirmed the expression of VE-cadherin in the differentiated cells by immunostaining (Fig. 1C). The above results held true regardless of whether the tissue was dermal or muscle tissue or of adipogenic origin. Taken together, our data strongly suggest that spheres represent a nonhematopoietic and nonendothelial subpopulation of cells associated with blood vessels growing from the heterogeneous Sca1+CD31–Lin– cell-sorted population of various adult tissues with the potential to differentiate into endothelial cells (summarized in Fig. 1D).

Developmental Origin of Tissue-Derived Spheres
Stem cells isolated from the dermis portion of craniofacial skin of juvenile mice (skin-derived progenitors [SKPs]) can be cultured as spheres and induced to differentiate in culture to produce neurons, glia, smooth muscle cells, and adipocytes [34]. Lineage analysis has indicated that both hair and whisker follicle dermal papillae contain neural crest-derived cells and that SKPs from the whisker pad are of neural crest origin [35]. To investigate whether the spheres we isolated from adult muscle, adipose, and dermis were also derived from the neural crest, we used mice that express cre recombinase under the control of a well-defined neural crest gene (Wnt-1) or endothelial-specific gene (Tie-2). Wnt-1-Cre and Tie-2-Cre mice were mated with R26R mice containing a β-galactosidase expression cassette preceded with a loxP-flanked stop sequence [36]. Cells expressing cre recombinase, as well as their descendants, will express β-galactosidase, as their loxP-flanked stop sequences would be excised. We found that spheres derived from the dermis (thoracic and abdominal), adipose, and muscle tissues were β-galactosidase+ when isolated from the Tie-2-Cre x R26R mice but not from the Wnt-1 x R26R mice (Fig. 1E). Therefore, these results unequivocally indicate that tissue-derived spheres from muscle, adipose, and dermis are not neural crest-derived, but rather are likely of mesodermal origin.

Differentiation Potential of Tissue-Derived Spheres
The differentiation potential of spheres derived from adipose, muscle, and dermal tissues was investigated and compared with eNSC that were also isolated as spheres. Cells cultured as spheres derived from muscle, adipose, or dermal tissue uniformly expressed nestin, a protein that is considered a marker of NSC (Fig. 2A; Table 2). Therefore, we investigated the ability of these cells to give rise to neural derivatives. In neural differentiation conditions, very small numbers of cells expressed neuronal (β-III-tubulin) or glial (GFAP) markers (Table 2). By contrast, the eNSC gave rise to a high proportion of cells expressing β-III-tubulin (94%) and GFAP (87%) (Table 2).


Figure 2
View larger version (51K):
[in this window]
[in a new window]

  		Figure 2. Spheres possess angiogenic characteristics in vitro. (A): Characterization of eNSC and spheres isolated from total cells of various tissues cultured in endothelial cell differentiation conditions. eNSC and spheres isolated from total cells of dermal, adipose, and muscle tissues were stained for nestin and CD31 and assessed for Ac-LDL uptake. All cell types expressed nestin, but only spheres expressed CD31 and incorporated Ac-LDL. (B): Flt-1/LacZ-derived eNSC and spheres from dermal, adipose, and muscle tissue grown on Matrigel and stained for β-galactosidase activity. On Matrigel, only spheres derived from Flt-1/LacZ mice formed capillary-like structures that were positive for β-galactosidase. (C): Semiquantitative reverse transcription-polymerase chain reaction for CD31, Flk-1, Tie-1, and G3PDH of 10T1/2-negative control cells, eNSC, and spheres isolated from total cells of dermal, adipose, and muscle tissues that were differentiated in Matrigel. Note that only spheres expressed endothelial-specific genes under differentiating conditions in Matrigel. Abbreviations: Ac-LDL, acetylated low-density lipoprotein; eNSC, embryo-derived neural stem cells; G3DPH, glyceraldehyde-3-phosphate dehydrogenase; rt, reverse transcriptase.

 


View this table:
[in this window]
[in a new window]

  		Table 2. Cells forming spheres are a subpopulation of resident tissue endothelial precursors

 
In adipogenic medium, tissue-derived spheres gave rise to a maximum of 8% adipocytes as scored by oil red O staining. By contrast, under the same conditions, eNSC never gave rise to adipocytes (Table 2). Similarly, under myogenic differentiation conditions, tissue-derived spheres gave rise only to rare cells that expressed desmin or MyoD, and no myotubes were ever detected. Therefore, we concluded that spheres derived from muscle, adipose, or dermis did not display robust neurogenic, adipogenic, or myogenic potential in vitro.

To test the angiogenic differentiation potential of tissue-derived spheres, cells were cultured in endothelial cell differentiation medium. Notably, cells dispersed from spheres uniformly did not express CD31. However after culture in differentiation-inducing conditions, high numbers of cells differentiated into apparently mature endothelial cells. Indeed, more than 90% of differentiated cells expressed high levels of CD31 (Fig. 2A; Table 2). By contrast, eNSC differentiated in the same manner displayed no detectable CD31 expression.

To further investigate the endothelial nature of the cells differentiated from tissue-derived spheres, we used a functional assay that assesses the ability of cells to take up acetylated low-density lipoprotein (Ac-LDL), an archetypal endothelial cell metabolic function (Table 2). We observed that approximately 98% of differentiated spheres incorporated high levels of fluorescent-tagged Ac-LDL, as compared with 47% for eNSC. Therefore, these data are consistent with the hypothesis that the differentiated spheres have an endothelial cell disposition.

To investigate angiogenic potential of spheres derived from muscle, adipose, and dermis, cells were cultured on Matrigel, a medium rich in basal lamina extracellular matrix proteins that promote endothelial cell differentiation and capillary formation [37, 38]. Under these growth conditions, tissue-derived spheres readily formed cord-like structures resembling angiogenic vessels, whereas the cultured eNSC were unable to do so. To determine the nature of the structures formed, Flt-1, an endothelial cell-specific receptor tyrosine kinase [39], was assessed by isolating and culturing spheres on Matrigel that were derived from Flt-1-lacZ knock-in mice [40]. Strikingly, the cord-like structures uniformly expressed high levels of β-galactosidase, confirming the angiogenic potential of the spheres (Fig. 2B).

The presence of endothelial precursors within spheres isolated from total cells of muscle, dermal, or adipose tissues was confirmed by semiquantitative PCR using specific primers to endothelial cell molecular markers CD31, Flk-1, and Tie-1 (Fig. 2C). Differentiation of the cells on Matrigel resulted in the expression of these angiogenic markers regardless of their tissue derivation. In contrast, eNSC and 10T1/2 cells did not express CD31, Flk-1, and Tie-1 when cultured on Matrigel (Fig. 2C). Taken together, these experiments strongly suggest that tissue-derived spheres are precursor cells with a robust potential to differentiate into endothelial cells. Therefore, we named these tissue-resident endothelial precursor cells.

Microarray Phenotyping Indicates That TEPs Are Highly Related in Distinct Tissues
To investigate the genetic phenotype of TEPs from different tissues, microarray analysis was conducted on TEPs isolated from muscle, adipose, and dermis. The resulting gene expression profiles were then compared with eNSC and other cells types. All gene expression data were deposited at StemBase (http://www.stembase.ca/; identifier E158). Hierarchical clustering of the hybridization values displayed a difference in expression patterns between the TEPs compared with eNSC (Fig. 3A). TEPs displayed a similar gene expression pattern, consistent with the hypothesis that they are related to one another and play a similar role within the three tissues.


Figure 3
View larger version (29K):
[in this window]
[in a new window]

  		Figure 3. TEPs, regardless of tissue derivation, display a similar molecular signature. (A): Heat map of the hybridization values obtained for the Affymetrix gene chip (MOE430A/B) probe sets of muscle-, adipose-, and dermal-derived TEP and eNSC preparations. Coloring indicates hybridization (high, red; low, green) in log scale, and intensity is related to the p-value of the measurement (with darker color for lower-reliability measurements). A standard hierarchical clustering was performed over all probe sets using the values of hybridization. Muscle, adipose and dermal samples clustered together. (B): Principal component analysis of the gene expression data. Gene expression values for 10T1/2 cells were used as a reference to subtract hybridization probes in common with the other probe sets. The graph displays the vector projections in the two directions given by the eigenvectors corresponding to the highest eigenvalues. Samples were eNSC, muscle, dermal, adipose, Meso A, and Meso D. The analysis shows that the expression signatures of the TEPs from the three tissues analyzed were similar to each other but different from the eNSC signature. (C): Venn diagram of probe sets that were specifically present in TEPs, eNSC, and 10T1/2 cells from a total of 10,231 positive probe sets. Abbreviations: eNSC, embryo-derived neural stem cells; Meso, mesoangioblast clone; TEP, tissue-resident endothelial precursor.

 
The similarity in gene expression for TEPs was further verified by principal component analysis of the hybridization patterns, including mesoangioblasts, an embryonic blood vessel-derived stem cell that readily differentiates into endothelial cells, as well as other mesodermal derivatives [41].

Microarray analysis was extended by subtracting genes that were not specific to TEPs. A Venn diagram of probe sets that are specifically hybridized in TEPs, eNSC, and 10T1/2 fibroblasts indicated that from a total of 10,231 positive probe sets, 833 were specific to TEPs (Fig. 3C; supplemental online Table 1). Taken together, gene expression analyses strongly supported the notion that TEPs represent a novel class of endothelial precursor cells. Twenty-five genes related to angiogenesis and endothelial cells were expressed in TEPs but not in eNSC (supplemental online Table 2). Therefore, the microarray analysis suggests that TEPs and eNSC are clearly two distinct cell types despite their isolation and growth as spheres.

TEPs Display Angiogenic Potential In Vivo
We first tested the differentiation potential of the cells that form TEPs in culture, the Sca1+/CD31–-sorted cellular fraction, in regenerating tissue in vivo. Freshly sorted Sca1+CD31– cells were isolated from dermis, adipose, or skeletal muscle tissue of Tg(GFPU)5Nagy/J mice that ubiquitously express GFP [42]. The GFP+Sca1+CD31– cells were injected into the regenerating TA muscle of a Balb/C mouse 4 days following injection with CTX, which induces the degeneration of muscle fibers [43]. Ten days post-transplantation, the recipient TA was recovered and isolated cells were analyzed by flow cytometry. Flow cytometric analysis revealed that between 40% and 70% of GFP+ cells had acquired CD31 expression, indicating that the transplanted cells had differentiated into mature endothelial cells regardless of tissue origin following injection (Fig. 4A). This suggests that the fraction of cells that give rise to TEPs can form a large number of endothelial cells, as well as other cell types.


Figure 4
View larger version (65K):
[in this window]
[in a new window]

  		Figure 4. Tissue-resident endothelial precursors (TEPs) possess angiogenic potential in vivo. (A): GFP+Sca1+CD31– cell fraction isolated from a GFP-Nagy muscle was transplanted into regenerating tibialis anterior (TA) muscle of a Balb/c mouse. Ten days postimplantation, muscle was digested and GFP+ cells were analyzed by flow cytometry. Results showed that a high percentage of donor GFP+Sca1+CD31– cells had converted to CD31+cells. (B): Regenerating TA S and WM 10 days post-transplantation with eNSC or Sca1+CD31–Lin– TEPs of dermal, muscle, and adipose origin from Flt-1/LacZ mice. Note that only TEPs formed capillary structures, as demonstrated by positive β-galactosidase staining regardless of tissue of origin. Abbreviations: APC, allophycocyanin; Comp, compensated; eNSC, embryo-derived neural stem cells; GFP, green fluorescent protein; PE, phycoerythrin; S, muscle section; SSC, side-scattered light; WM, whole mount.

 
We next investigated the capacity of dermis-, adipose-, or muscle tissue-derived Sca1+CD31–Lin– TEPs isolated from Flt-1-LacZ mice to differentiate into endothelial cells in vivo. Flt-1 activity was assessed by β-galactosidase staining 10 days after TEP implantation into CTX-damaged muscle. Strikingly, we observed extensive networks of Flt-1-LacZ blood vessels spread throughout the regenerated muscle, regardless of the tissue of origin of the TEPs (Fig. 4B). By contrast, eNSC isolated from Flt-1-LacZ embryos did not exhibit migration or angiogenic activity. Taken together, our data confirm that TEPs are endothelial precursors that readily participate in neovascularization during tissue regeneration.

To investigate the in vivo relevance of TEPs to neovascularization during muscle regeneration, the abundance of TEPs was assessed by a sphere-forming assay 4 days after CTX injection. We observed a 13.6-fold increase in the total number of mononuclear cells per TA, due in part to the presence of inflammatory cells at the site of injury (Fig. 5A). For TEP enumeration, the inflammatory cells within our cellular fractions were removed by isolating the Sca1+CD31–Lin– population. We observed a marked increase in the fraction of Sca1+CD31–Lin– cells containing TEPs; notably, 25.3% of the total number of cells were Sca1+CD31–Lin–, as opposed to 7.2% in noninjured muscle (Fig. 5B).


Figure 5
View larger version (91K):
[in this window]
[in a new window]

  		Figure 5. Transplanted TEPs contribute to vasculogenesis during muscle regeneration. (A): Graphical representation of isolated cells per mg of tibialis anterior (TA) that was noninsulted (control) or CTX-insulted post-4 days. Asterisk denotes significance (p < .001). (B): Graphical representation of the percentage of cells from (A) that were Sca1+CD31–Lin–. Asterisk denotes significance (p < .001). (C): Graphical representation of the number of TEPs per mg TA from noninsulted (control) or CTX-insulted after 4 days. Asterisk denotes significance (p < .001). (D): Cultured TEPs from TA of noninsulted (control) and CTX-insulted post-4 days under growth and differentiation (Matrigel) conditions. Capillary formation was visualized by staining for alkaline phosphatase. Abbreviations: CTX, cardiotoxin; Nb., number; TEP, tissue-resident endothelial precursor.

 
To quantify the relative increase in numbers of TEPs that were present within the Sca1+CD31–Lin– cell fraction, equal cell numbers were plated on a methylcellulose-based medium that supports growth of spheres. After 10 days in culture, spheres were present in a ratio of 1 per 5,507 cells from noninjured muscle as compared with 1 per 574 cells from CTX-injured muscle. The total number of TEPs per gram of muscle was almost 48-fold higher in regenerating muscle 4 days after CTX injection (Fig. 5C). Furthermore, spheres from normal and CTX-injured muscle, when plated on Matrigel, both gave rise to capillary-like structures that stained positively for ALKP, a marker of endothelial cells (Fig. 5D).

Taken together, the above data strongly suggest that TEPs directly participate in neovascularization during the muscle regeneration. Moreover, TEPs derived from uninjured or regenerating muscle displayed the same angiogenic potential. Therefore, TEPs appear to represent an adult mouse tissue-resident endothelial precursor that actively participates in neovascularization during tissue regeneration. The establishment of their exact role during the kinetics of vascular remodeling or capillary formation in vivo remains to be detailed.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 Acknowledgments
 References
 
Blood vessels supply nutrients and oxygen to all tissues, allowing them to function efficiently. In normal conditions, cells lining blood vessels display a low turnover, their replacement contingent on the division and migration of adjacent mature endothelial cells [4, 44]. This phenomenon is accentuated during vasculogenesis, when a rapid recruitment of endothelial cells is required for adequate neovascularization of the tissues. Our results describe the isolation and function of an adult mouse resident tissue endothelial precursor/stem cell that is involved in this process. We present evidence for the existence of adult resident endothelial precursors, TEPs, which grow as spheres in culture. Our findings now add a third distinct population of endothelial progenitor cells, along with bone marrow-derived EPC and monocyte/macrophage cells, that are capable of repairing damaged vessels. There is much evidence for the role of TEPs in neovascularization. Microarray analyses show that TEPs possess similar molecular signatures regardless of their tissue derivation, expressing genes related to angiogenesis/endothelial cells. In vitro and in vivo differentiation assays have revealed that TEPs readily differentiate into endothelial cells and thus function as endothelial precursors. However, the relationship of TEPs to the well-characterized endothelial progenitor cells [45, 46] remains to be established. In addition, TEPs are associated with blood vessels, a perfect location for recruitment and maintenance of the vasculature during severe trauma when neovascularization is required. Their physiological importance is demonstrated by their expansion in vivo following a chemically induced muscle injury and subsequent contribution to the vasculature of regenerating tissues.

Before our report, the biological contribution of stem/precursor endothelial cells during tissue regeneration had not been sufficiently characterized. Studies had relied only on transplantation to confirm the potential of spheres to give rise to donor-derived endothelial cells as part of the recipient's reconstructed vasculature [18, 47, 48]. Other studies had used direct intramuscular injection of EPC or BM transplantation of genetically engineered EPC [8, 49–51] (reviewed by Urbich and Dimmeler [3]). Although the physiological contribution of EC progenitors in vivo was not quantified in these studies, the approaches shed light on the mechanisms required for homing of EPC to sites of inflammation and their subsequent differentiation [52, 53].

We found that TEPs derived from resting muscle have the same angiogenic properties as regenerating muscle after induced trauma, suggesting that they might also be involved in maintaining the vasculature under normal physiological conditions. This is unlike the recruitment of EPC, which are apparent only during tissue trauma or pathologic states.

Although proinflammatory cytokines such as tumor necrosis factor regulate the expression of Sca1 in T and B cells, we are confident that the increase in Sca1+ cells that we observed during wound healing is not biased by inflammatory cells [54, 55]. Indeed, our experiment setup showed that TEPs grown from the Sca1+CD31– were not derived from peripheral blood, since they were hematopoietic lineage-depleted (Lin–) or leukocyte-free. In addition, RT-PCR analyses demonstrated a lack of CD45 expression in the freshly sorted fractions. Finally, we were unable to grow TEPs from the blood-derived enriched mononuclear cell fraction or from Lin+ cells (Table 1).

Our differentiation assays strongly suggest that TEPs are adult resident angiogenic progenitors, in agreement with previously published data, rather than multipotent cells as earlier reported [18, 34, 47, 48]. Indeed, only a low percentage of adipocytes, neurons, and myocytes were obtained under various culture conditions, as opposed to the substantial rate of endothelial differentiation. In addition, nestin expression does not represent a neuronal progenitor marker, since it has been described in a variety of other cell types, including skeletal muscle and endothelial cells [56–62]. Ultimately, RT-PCR examination and immunostaining of TEPs grown in endothelial-specific culture conditions clearly show the expression of various endothelial cell markers, including VE-cadherin.

β-Galactosidase-positive expression of TEPs from Tie-2/R26R mice suggests their mesodermal origin rather than a neural crest origin as has been reported for other adult cells growing as spheres, such as SKPs and NSCs that were isolated from craniofacial skin and the central nervous system, respectively, both derived from neural crest [35, 63]. One of the major differences between TEPs and these neural crest-derived progenitor cells resides in their capacity to self-renew. Indeed, adult neural crest-derived cells have the potential to grow and to maintain long-term nondifferentiated cells for more than 50 passages [34]. In our case, we were unable to passage TEPs for more than five passages. Our results are in accordance with those of Gingras et al. that described adult spheres originating from thoracic skin with a very low proliferative index [64]. This technical aspect makes a clonal multipotency analysis of TEPs extremely difficult. Moreover, the fact that TEPs are unable to proliferate efficiently in vitro corroborates numerous reports describing murine-derived endothelial cells that can grow in vitro only when retrovirally infected with polyoma middle T antigen [65]. Therefore, in the hierarchy of stem cells, TEPs may represent committed cells with a lower proliferative index and self-renewal capacity, and/or factors that allow for their proliferation in vitro may be suboptimal. In future experiments, it would be of interest to develop a murine system to investigate the relationship between TEPs and other cells with endothelial potential, such as endothelial progenitors, as has been performed by Ingram et al. for human endothelial progenitor cells [45, 46].

TEPs may originate from primitive progenitors during development. Indeed, recent studies have demonstrated the existence of the hemangioblast, a precursor cell with the potential to differentiate into both whole blood cells and blood vessels consisting of endothelial and smooth muscle cells during embryogenesis [1, 66]. Such primitive progenitor cells stay in muscle tissue until adulthood, required for maintenance and repair. In addition, previous studies have established that the limb vasculature derived from somites and that their dorsolateral regions are a source of vessel endothelia in limbs [67, 68]. Kardon et al. analyzed the somatic-derived cell lineages in the chicken embryo and found that most somite-derived cells were bipotential and could give rise to both muscle and endothelial cells [69]. Indeed, CD34+Flk1+ endothelial progenitor cells from mouse fetal limb muscle gave rise to myogenic cells in vitro and when transplanted into adult mouse muscle [70]. However, the migration of this fetal cell population into dermal and adipose tissues, where TEPs also reside, has not yet been demonstrated. More importantly, we were unable to differentiate TEPs into myocytes in vitro and in vivo (data not shown), suggesting that if TEPs were somite-derived cells, they did not preserve their myogenic potential. However, this does not rule out the possibility that TEPs may originate from a committed angioblast derived from a hemangioblast, as reported by the Cossu laboratory, that can give rise to muscle and endothelial cells [41, 71, 72].

In conclusion, we have characterized a novel class of ubiquitous mouse adult resident angiogenic precursors found on small blood vessels of various tissues. These cells were grown as spheres in vitro and participated in capillary vessel formation when transplanted into regenerating muscle. In vivo, their numbers were shown to significantly increase concomitant to their participation in the wound healing process. It will be advantageous to verify their exact contribution to this process. In addition, possible interactions with EPC and monocytic-derived endothelial progenitors in mouse and human tissue following trauma are yet to be determined. The study of TEPs is of interest in evaluating their potential contribution during cancer angiogenesis and for an autonomous source of precursor cells for cell therapy, vascular prosthesis endothelialization, and tissue engineering. Furthermore, resident cells may provide another cell source to stimulate neovascularization and collateral blood vessel formation when required during pathological disturbances, such as atherosclerosis.


   DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
Acknowledgments
 References
 
The authors indicate no potential conflicts of interest.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
References
 
We thank Dr. Iain McKinnell for critical reading of the manuscript; Caroline Vergette for cell sorting assistance; Dr. Stefano Ferrari for access to his microarray database; and Dr. Francois Berthod and Marie-France Champigny for insight into isolation of TEPs. We gratefully acknowledge Dr. Guo-Hua Fong for providing us Flt-1/LacZ transgenic mice. G.G. was a recipient of a postdoctoral fellowship from the Fonds de la Recherche en Santé du Québec. M.A.A.-N. is a recipient of a Canada Research Chair in Bioinformatics. This work was supported by grants from the Canadian Institutes of Health Research. M.A.R. is an International Scholar of the Howard Hughes Medical Institute and holds the Canada Research Chair in Molecular Genetics. G.G. is currently affiliated with Department of Orthopaedic Surgery, Faculty of Medicine, Research Center on Aging, Sherbrooke, Québec, Canada. A.A. is currently affiliated with University of Minnesota Medical School, Minneapolis, MN.


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 

  1. Ema M, Rossant J. Cell fate decisions in early blood vessel formation. Trends Cardiovasc Med 2003;13:254–259.[CrossRef][Medline]

  2. Folkman J. Angiogenesis in cancer, vascular, rheumatoid and other diseases. Nat Med 1995;1:27–31.[CrossRef][Medline]

  3. Urbich C, Dimmeler S. Endothelial progenitor cells functional characterization. Trends Cardiovasc Med 2004;14:318–322.[CrossRef][Medline]

  4. Wagner EF, Risau W. Oncogenes in the study of endothelial cell growth and differentiation. Semin Cancer Biol 1994;5:137–145.[Medline]

  5. Hristov M, Erl W, Weber PC. Endothelial progenitor cells: Mobilization, differentiation, and homing. Arterioscler Thromb Vasc Biol 2003;23:1185–1189.[Abstract/Free Full Text]

  6. Asahara T, Murohara T, Sullivan A et al. Isolation of putative progenitor endothelial cells for angiogenesis. Science 1997;275:964–967.[Abstract/Free Full Text]

  7. Takahashi T, Kalka C, Masuda H et al. Ischemia- and cytokine-induced mobilization of bone marrow-derived endothelial progenitor cells for neovascularization. Nat Med 1999;5:434–438.[CrossRef][Medline]

  8. Asahara T, Masuda H, Takahashi T et al. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res 1999;85:221–228.[Abstract/Free Full Text]

  9. Biancone L, Cantaluppi V, Duo D et al. Role of L-selectin in the vascular homing of peripheral blood-derived endothelial progenitor cells. J Immunol 2004;173:5268–5274.[Abstract/Free Full Text]

  10. Anghelina M, Moldovan L, Zabuawala T et al. A subpopulation of peritoneal macrophages form capillary-like lumens and branching patterns in vitro. J Cell Mol Med 2006;10:708–715.[Medline]

  11. Nowak G, Karrar A, Holmen C et al. Expression of vascular endothelial growth factor receptor-2 or Tie-2 on peripheral blood cells defines functionally competent cell populations capable of reendothelialization. Circulation 2004;110:3699–3707.[Abstract/Free Full Text]

  12. Rehman J, Li J, Orschell CM et al. Peripheral blood "endothelial progenitor cells" are derived from monocyte/macrophages and secrete angiogenic growth factors. Circulation 2003;107:1164–1169.[Abstract/Free Full Text]

  13. Schmeisser A, Garlichs CD, Zhang H et al. Monocytes coexpress endothelial and macrophagocytic lineage markers and form cord-like structures in Matrigel under angiogenic conditions. Cardiovasc Res 2001;49:671–680.[Abstract/Free Full Text]

  14. Conejo-Garcia JR, Buckanovich RJ, Benencia F et al. Vascular leukocytes contribute to tumor vascularization. Blood 2005;105:679–681.[Abstract/Free Full Text]

  15. Urbich C, Heeschen C, Aicher A et al. Relevance of monocytic features for neovascularization capacity of circulating endothelial progenitor cells. Circulation 2003;108:2511–2516.[Abstract/Free Full Text]

  16. Miranville A, Heeschen C, Sengenes C et al. Improvement of postnatal neovascularization by human adipose tissue-derived stem cells. Circulation 2004;110:349–355.[Abstract/Free Full Text]

  17. Planat-Benard V, Silvestre JS, Cousin B et al. Plasticity of human adipose lineage cells toward endothelial cells: Physiological and therapeutic perspectives. Circulation 2004;109:656–663.[Abstract/Free Full Text]

  18. Beltrami AP, Barlucchi L, Torella D et al. Adult cardiac stem cells are multipotent and support myocardial regeneration. Cell 2003;114:763–776.[CrossRef][Medline]

  19. Wurmser AE, Nakashima K, Summers RG et al. Cell fusion-independent differentiation of neural stem cells to the endothelial lineage. Nature 2004;430:350–356.[CrossRef][Medline]

  20. Majka SM, Jackson KA, Kienstra KA et al. 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]

  21. Gussoni E, Soneoka Y, Strickland CD et al. Dystrophin expression in the mdx mouse restored by stem cell transplantation. Nature 1999;401:390–394.[CrossRef][Medline]

  22. Asakura A, Seale P, Girgis-Gabardo A et al. Myogenic specification of side population cells in skeletal muscle. J Cell Biol 2002;159:123–134.[Abstract/Free Full Text]

  23. Tamaki T, Akatsuka A, Ando K et al. Identification of myogenic-endothelial progenitor cells in the interstitial spaces of skeletal muscle. J Cell Biol 2002;157:571–577.[Abstract/Free Full Text]

  24. Qu-Petersen Z, Deasy B, Jankowski R et al. Identification of a novel population of muscle stem cells in mice: Potential for muscle regeneration. J Cell Biol 2002;157:851–864.[Abstract/Free Full Text]

  25. Kablar B, Krastel K, Ying C et al. MyoD and Myf-5 differentially regulate the development of limb versus trunk skeletal muscle. Development 1997;124:4729–4738.[Abstract]

  26. Xie Y, Muller WA. Molecular cloning and adhesive properties of murine platelet/endothelial cell adhesion molecule 1. Proc Natl Acad Sci U S A 1993;90:5569–5573.[Abstract/Free Full Text]

  27. Vittet D, Prandini MH, Berthier R et al. Embryonic stem cells differentiate in vitro to endothelial cells through successive maturation steps. Blood 1996;88:3424–3431.[Abstract/Free Full Text]

  28. Oelrichs RB, Reid HH, Bernard O et al. NYK/FLK-1: A putative receptor protein tyrosine kinase isolated from E10 embryonic neuroepithelium is expressed in endothelial cells of the developing embryo. Oncogene 1993;8:11–18.[Medline]

  29. Asashima T, Iizasa H, Terasaki T et al. Rat brain pericyte cell lines expressing beta2-adrenergic receptor, angiotensin II receptor type 1A, klotho, and CXCR4 mRNAs despite having endothelial cell markers. J Cell Physiol 2003;197:69–76.[CrossRef][Medline]

  30. Asakura A, Komaki M, Rudnicki M. Muscle satellite cells are multipotential stem cells that exhibit myogenic, osteogenic, and adipogenic differentiation. Differentiation 2001;68:245–253.[CrossRef][Medline]

  31. Weiss S, Dunne C, Hewson J et al. Multipotent CNS stem cells are present in the adult mammalian spinal cord and ventricular neuroaxis. J Neurosci 1996;16:7599–7609.[Abstract/Free Full Text]

  32. Reynolds BA, Tetzlaff W, Weiss S. A multipotent EGF-responsive striatal embryonic progenitor cell produces neurons and astrocytes. J Neurosci 1992;12:4565–4574.[Abstract]

  33. van de Rijn M, Heimfeld S, Spangrude GJ et al. Mouse hematopoietic stem-cell antigen Sca-1 is a member of the Ly-6 antigen family. Proc Natl Acad Sci U S A 1989;86:4634–4638.[Abstract/Free Full Text]

  34. Toma JG, Akhavan M, Fernandes KJ et al. Isolation of multipotent adult stem cells from the dermis of mammalian skin. Nat Cell Biol 2001;3:778–784.[CrossRef][Medline]

  35. Fernandes KJ, McKenzie IA, Mill P et al. A dermal niche for multipotent adult skin-derived precursor cells. Nat Cell Biol 2004;6:1082–1093.[CrossRef][Medline]

  36. Chai Y, Jiang X, Ito Y et al. Fate of the mammalian cranial neural crest during tooth and mandibular morphogenesis. Development 2000;127:1671–1679.[Abstract]

  37. Grant DS, Tashiro K, Segui-Real B et al. Two different laminin domains mediate the differentiation of human endothelial cells into capillary-like structures in vitro. Cell 1989;58:933–943.[CrossRef][Medline]

  38. Kubota Y, Kleinman HK, Martin GR et al. Role of laminin and basement membrane in the morphological differentiation of human endothelial cells into capillary-like structures. J Cell Biol 1988;107:1589–1598.[Abstract/Free Full Text]

  39. He Y, Smith SK, Day KA et al. Alternative splicing of vascular endothelial growth factor (VEGF)-R1 (FLT-1) pre-mRNA is important for the regulation of VEGF activity. Mol Endocrinol 1999;13:537–545.[Abstract/Free Full Text]

  40. Fong GH, Rossant J, Gertsenstein M et al. Role of the Flt-1 receptor tyrosine kinase in regulating the assembly of vascular endothelium. Nature 1995;376:66–70.[CrossRef][Medline]

  41. Minasi MG, Riminucci M, De Angelis L et al. The meso-angioblast: A multipotent, self-renewing cell that originates from the dorsal aorta and differentiates into most mesodermal tissues. Development 2002;129:2773–2783.[Abstract/Free Full Text]

  42. Hadjantonakis AK, Gertsenstein M, Ikawa M et al. Generating green fluorescent mice by germline transmission of green fluorescent ES cells. Mech Dev 1998;76:79–90.[CrossRef][Medline]

  43. Charge SB, Rudnicki MA. Cellular and molecular regulation of muscle regeneration. Physiol Rev 2004;84:209–238.[Abstract/Free Full Text]

  44. Werner N, Nickenig G. Clinical and therapeutical implications of EPC biology in atherosclerosis. J Cell Mol Med 2006;10:318–332.[Medline]

  45. Ingram DA, Mead LE, Moore DB et al. Vessel wall-derived endothelial cells rapidly proliferate because they contain a complete hierarchy of endothelial progenitor cells. Blood 2005;105:2783–2786.[Abstract/Free Full Text]

  46. Ingram DA, Mead LE, Tanaka H et al. Identification of a novel hierarchy of endothelial progenitor cells using human peripheral and umbilical cord blood. Blood 2004;104:2752–2760.[Abstract/Free Full Text]

  47. Belicchi M, Pisati F, Lopa R et al. Human skin-derived stem cells migrate throughout forebrain and differentiate into astrocytes after injection into adult mouse brain. J Neurosci Res 2004;77:475–486.[CrossRef][Medline]

  48. Gorio A, Torrente Y, Madaschi L et al. Fate of autologous dermal stem cells transplanted into the spinal cord after traumatic injury (TSCI). Neuroscience 2004;125:179–189.[CrossRef][Medline]

  49. Kocher AA, Schuster MD, Szabolcs MJ et al. Neovascularization of ischemic myocardium by human bone-marrow-derived angioblasts prevents cardiomyocyte apoptosis, reduces remodeling and improves cardiac function. Nat Med 2001;7:430–436.[CrossRef][Medline]

  50. Carmeliet P, Luttun A. The emerging role of the bone marrow-derived stem cells in (therapeutic) angiogenesis. Thromb Haemost 2001;86:289–297.[Medline]

  51. Friedrich EB, Walenta K, Scharlau J et al. CD34-/CD133+/VEGFR-2+ endothelial progenitor cell subpopulation with potent vasoregenerative capacities. Circ Res 2006;98:e20–25.[Abstract/Free Full Text]

  52. Masuda H, Asahara T. Post-natal endothelial progenitor cells for neovascularization in tissue regeneration. Cardiovasc Res 2003;58:390–398.[Abstract/Free Full Text]

  53. Rabbany SY, Heissig B, Hattori K et al. Molecular pathways regulating mobilization of marrow-derived stem cells for tissue revascularization. Trends Mol Med 2003;9:109–117.[CrossRef][Medline]

  54. Luna G, Paez J, Cardier JE. Expression of the hematopoietic stem cell antigen Sca-1 (LY-6A/E) in liver sinusoidal endothelial cells: Possible function of Sca-1 in endothelial cells. Stem Cells Dev 2004;13:528–535.[CrossRef][Medline]

  55. Malek TR, Danis KM, Codias EK. Tumor necrosis factor synergistically acts with IFN-gamma to regulate Ly-6A/E expression in T lymphocytes, thymocytes and bone marrow cells. J Immunol 1989;142:1929–1936.[Abstract]

  56. Vaittinen S, Lukka R, Sahlgren C et al. The expression of intermediate filament protein nestin as related to vimentin and desmin in regenerating skeletal muscle. J Neuropathol Exp Neurol 2001;60:588–597.[Medline]

  57. Sejersen T, Lendahl U. Transient expression of the intermediate filament nestin during skeletal muscle development. J Cell Sci 1993;106:1291–1300.[Abstract]

  58. Mokry J, Nemecek S. Cerebral angiogenesis shows nestin expression in endothelial cells. Gen Physiol Biophys 1999;18 (suppl 1):25–29.[Medline]

  59. Mokr J, Nemecek S. Angiogenesis of extra- and intraembryonic blood vessels is associated with expression of nestin in endothelial cells. Folia Biol (Praha) 1998;44:155–161.[Medline]

  60. Klein T, Ling Z, Heimberg H et al. Nestin is expressed in vascular endothelial cells in the adult human pancreas. J Histochem Cytochem 2003;51:697–706.[Abstract/Free Full Text]

  61. Lardon J, Rooman I, Bouwens L. Nestin expression in pancreatic stellate cells and angiogenic endothelial cells. Histochem Cell Biol 2002;117:535–540.[CrossRef][Medline]

  62. Mokry J, Nemecek S. Immunohistochemical detection of intermediate filament nestin. Acta Medica (Hradec Kralove) 1998;41:73–80.

  63. Rao M. Stem and precursor cells in the nervous system. J Neurotrauma 2004;21:415–427.[CrossRef][Medline]

  64. Gingras M, Champigny MF, Berthod F. Differentiation of human adult skin-derived neuronal precursors into mature neurons. J Cell Physiol 2007;210:498–506.[CrossRef][Medline]

  65. Pepper MS, Tacchini-Cottier F, Sabapathy TK et al. Endothelial cells transformed by polyoma virus middle T oncogene: A model for hemangiomas and other vascular tumors. In: Bicknell R, Lewis CE, Ferrara N, eds. Tumour Angiogenesis.Oxford, U.K.: Oxford University Press,1997;309–331.

  66. Huber TL, Kouskoff V, Fehling HJ et al. Haemangioblast commitment is initiated in the primitive streak of the mouse embryo. Nature 2004;432:625–630.[CrossRef][Medline]

  67. Wilting J, Brand-Saberi B, Huang R et al. Angiogenic potential of the avian somite. Dev Dyn 1995;202:165–171.[Medline]

  68. Buckingham M, Bajard L, Chang T et al. The formation of skeletal muscle: From somite to limb. J Anat 2003;202:59–68.[CrossRef][Medline]

  69. Kardon G, Campbell JK, Tabin CJ. Local extrinsic signals determine muscle and endothelial cell fate and patterning in the vertebrate limb. Dev Cell 2002;3:533–545.[CrossRef][Medline]

  70. Le Grand F, Auda-Boucher G, Levitsky D et al. Endothelial cells within embryonic skeletal muscles: A potential source of myogenic progenitors. Exp Cell Res 2004;301:232–241.[CrossRef][Medline]

  71. De Angelis L, Berghella L, Coletta M et al. Skeletal myogenic progenitors originating from embryonic dorsal aorta coexpress endothelial and myogenic markers and contribute to postnatal muscle growth and regeneration. J Cell Biol 1999;147:869–878.[Abstract/Free Full Text]

  72. Esner M, Meilhac SM, Relaix F et al. Smooth muscle of the dorsal aorta shares a common clonal origin with skeletal muscle of the myotome. Development 2006;133:737–749.[Abstract/Free Full Text]




This article has been cited by other articles:


Home page
 Cancer Res.Home page
Y. Zhang, A. Daquinag, D. O. Traktuev, F. Amaya-Manzanares, P. J. Simmons, K. L. March, R. Pasqualini, W. Arap, and M. G. Kolonin
White Adipose Tissue Cells Are Recruited by Experimental Tumors and Promote Cancer Progression in Mouse Models
Cancer Res., June 15, 2009; 69(12): 5259 - 5266.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
A. Z. Caron, G. Drouin, J. Desrosiers, F. Trensz, and G. Grenier
A novel hindlimb immobilization procedure for studying skeletal muscle atrophy and recovery in mouse
J Appl Physiol, June 1, 2009; 106(6): 2049 - 2059.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
2006-0795v1
2006-0795v2
25/12/3101    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Reprints/Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Grenier, G.
Right arrow Articles by Rudnicki, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Grenier, G.
Right arrow Articles by Rudnicki, M. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
STEM CELLS THE ONCOLOGIST CME ALPHAMED PRESS JOURNALS