The postnatal rat aorta contains pericyte progenitor cells that form spheroidal colonies in suspension culture

K. M. Howson,1 A. C. Aplin,1 M. Gelati,1,3 G. Alessandri,3 E. A. Parati,3 and R. F. Nicosia1,2

1Department of Pathology, University of Washington; 2Division of Pathology and Laboratory Medicine, Veterans Administration Puget Sound Health Care System, Seattle, WA; and 3Laboratory of Neurobiology and Neuroregenerative Therapy, "Carlo Besta" Institute, Milan, Italy

Submitted 21 April 2005 ; accepted in final form 28 July 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Pericytes play an important role in modulating angiogenesis, but the origin of these cells is poorly understood. To evaluate whether the mature vessel wall contains pericyte progenitor cells, nonendothelial mesenchymal cells isolated from the rat aorta were cultured in a serum-free medium optimized for stem cells. This method led to the isolation of anchorage-independent cells that proliferated slowly in suspension, forming spheroidal colonies. This process required basic fibroblast growth factor (bFGF) in the culture medium, because bFGF withdrawal caused the cells to attach to the culture dish and irreversibly lose their capacity to grow in suspension. Immunocytochemistry and RT-PCR analysis revealed the expression of the precursor cell markers CD34 and Tie-2 and the absence of endothelial cell markers (CD31 and endothelial nitric oxide synthase, eNOS) and smooth muscle cell markers ({alpha}-smooth muscle actin, {alpha}-SMA). In addition, spheroid-forming cells were positive for NG2, nestin, PDGF receptor (PDGFR)-{alpha}, and PDGFR-{beta}. Upon exposure to serum, these cells lost CD34 expression, acquired {alpha}-SMA, and attached to the culture dish. Returning these cells to serum-free medium failed to restore their original spheroid phenotype, suggesting terminal differentiation. When embedded in collagen gels, spheroid-forming cells rapidly migrated in response to PDGF-BB and became dendritic. Spheroid-forming cells cocultured in collagen with angiogenic outgrowths of rat aorta or isolated endothelial cells transformed into pericytes. These results demonstrate that the rat aorta contains primitive mesenchymal cells capable of pericyte differentiation. These immature cells may represent an important source of pericytes during angiogenesis in physiological and pathological processes. They may also provide a convenient supply of mural cells for vascular bioengineering applications.

angiogenesis; stem cells; smooth muscle; mural cells; collagen


BLOOD VESSELS FORM IN RESPONSE to angiogenic factor stimulation by two main processes: vasculogenesis and angiogenesis. Vasculogenesis is the de novo formation of vessels from immature mesenchyme, whereas angiogenesis refers to the sprouting of neovessels from a preexisting vasculature. Both vasculogenesis and angiogenesis occur in the developing embryo, whereas angiogenic sprouting is the main mechanism of neovascularization during postnatal wound healing, inflammation, and the female reproductive cycle (11, 48). Angiogenesis also contributes to the progression of many pathological conditions, including hemangiomas, cancer, atherosclerosis, proliferative retinopathy, rheumatoid arthritis, and psoriasis (6, 12).

Angiogenic sprouting in the adult was initially considered an exclusive feature of terminally differentiated endothelial cells, which have the capacity to form vascular tubes and recruit mural cells (pericytes or smooth muscle cells). Later studies, however, demonstrated that the angiogenic process might also involve bone marrow-derived vascular progenitor cells (3, 44). These immature cells circulate in the peripheral blood and colonize angiogenic sites, where they differentiate into endothelial cells or mural cells, becoming incorporated into the developing neovasculature (10, 46). Immature mesenchymal cells with vascular progenitor features have also been shown to reside in peripheral tissues such as skeletal muscle (30), where they participate in the angiogenic response to injury.

Studies conducted at our laboratory and by others have demonstrated that angiogenesis can be reproduced ex vivo by culturing explants of adult rat aorta or other blood vessel types in three-dimensional (3D) biomatrices (24, 36, 38, 39). Neovessels formed in this model are composed of a luminal layer of endothelial cells and surrounding pericytes. The origin of these neovessels has been attributed to differentiated cells of the vessel wall. However, there is evidence from experiments with fetal aortas that neovessels may also derive from primitive CD34-positive/CD31-negative cells (1), raising the possibility that immature mesenchymal cells contribute to the postnatal angiogenic response of the vessel wall. Despite the many studies of circulating vascular progenitor cells that have been performed, the involvement in postnatal angiogenesis of progenitor cells that reside in the vessel wall has not been evaluated.

The purpose of this study was to investigate the hypothesis that the postnatal aorta contains vascular progenitor cells and to develop culture conditions for their isolation and long-term maintenance in an undifferentiated state. To identify and isolate these cells, we adapted a method originally described for neural stem cells (42, 47). Isolated cells were characterized by performing immunofluorescent staining, RT-PCR, and electron microscopy as well as functional studies with 3D models of angiogenesis (35, 59).

In this report, we describe how morphologically primitive, CD34-positive/{alpha}-smooth muscle actin ({alpha}-SMA)-negative nonendothelial cells can be isolated from the mature rat aorta and propagated in suspension culture, where they grow as spheroid structures. These cells express markers of early pericyte lineage, can be induced to differentiate into CD34-negative/{alpha}-SMA-positive adherent cells, and behave functionally as pericytes when cocultured in collagen gels with vasoformative endothelial outgrowths. These immature mesenchymal cells may be an important source of mural cells in physiological or pathological angiogenic responses of the aortic wall.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Isolation and culture of pericyte progenitor cells. All animal procedures were performed in accordance with Veterans Administration Puget Sound Health Care System Institutional Animal Care and Use Committee and National Institutes of Health guidelines. Thoracic aortas were dissected from euthanized Fischer 344 male rats (Harlan, Indianapolis, IN). After excision, aortas were cleaned of fibroadipose tissue and blood, minced, and incubated for 3 h at 37°C in endothelial basal medium (EBM; Cambrex BioScience, Walkersville, MD) containing 0.25% collagenase D (Roche Applied Science, Indianapolis, IN) and 0.25% BSA (Invitrogen, Carlsbad, CA). The digest was filtered (10 µm) to obtain a single-cell suspension. CD31-positive cells were selectively removed with magnetic beads (CELLection Pan Mouse IgG kit; Dynal Biotech, Oslo, Norway) coated with anti-CD31 antibody (BD Biosciences Pharmingen, San Diego, CA). CD31-negative cells were transferred to a 30-mm tissue culture dish coated with 125 µg/ml rat tail collagen (9). Cells were cultured in H medium (42, 47) composed of DMEM/Ham's F-12 medium, 0.1125% NaHCO3, 2 mM glutamine, 0.4% BSA fraction V, 5 mM HEPES, 20 ng/ml recombinant human EGF (Invitrogen), 0.6% glucose, 4 µg/ml heparin, 100 µg/ml apotransferrin, 25 µg/ml insulin, 9.66 µg/ml putrescine, 20 µM progesterone, 30 µM selenium (Sigma-Aldrich, St. Louis, MO), 10 ng/ml recombinant bovine basic fibroblast growth factor (bFGF; Calbiochem, San Diego, CA), and 1 ng/ml recombinant human leukemia-inhibitory factor (LIF; Chemicon International, Temecula, CA). After 5–7 days, the nonadherent cells, named spheroid-forming aortic cells (SFACs) because of their pattern of growth in suspension, were transferred to an uncoated dish and maintained in a humidified incubator at 37°C in a 5% CO2 atmosphere. To induce differentiation, SFACs were cultured in H medium containing 10% FBS (Invitrogen). To study the migratory response of SFACs in a 3D matrix, cells were suspended in rat tail collagen (1 mg/ml) and grown in EBM containing 50 ng/ml of either VEGF, bFGF, or PDGF-BB (R&D Systems, Minneapolis, MN). Cultures containing no growth factors were used as controls.

Isolation and culture of rat aortic endothelial cells. Rat aortic endothelial cells (RAECs) were isolated from collagenase-digested aortic tissue, using anti-CD31 antibody-coated magnetic beads. CD31-positive RAECs were plated onto dishes coated with collagen and fibronectin (1 µg/cm2; Invitrogen). Cells were grown in EBM containing 10% FBS, 10 ng/ml EGF, 20 ng/ml bFGF, 10 ng/ml VEGF, and 10 µg/ml heparin.

Coculture of pericyte progenitor cells with aortic rings. The aortic ring assay of angiogenesis (59) was modified to include SFACs. Briefly, individual aortic rings were embedded in a 40-µl drop of 1 mg/ml rat tail collagen containing ~30 spheroids. After collagen gelation, cultures were grown in EBM containing 10 ng/ml VEGF. Pericyte transformation of SFACs was evaluated using phase-contrast microscopy and immunocytochemistry, followed by confocal microscopy.

Coculture of pericyte progenitor cells with endothelial cells. A capillary tube formation assay (33, 37, 52) was used to study interactions between SFACs and isolated RAECs in a 3D collagen matrix. SFACs and RAECs were suspended in rat tail collagen (4 x 105 of each cell type/ml) and plated in 16-mm wells (200 µl/well). After collagen gelation, cultures were incubated in EBM containing 10% FBS, 10 ng/ml EGF, 50 ng/ml bFGF, 50 ng/ml VEGF, and 10 µg/ml heparin. Control cultures consisted of RAECs embedded in collagen gels without SFACs. Pericyte transformation of SFACs was evaluated as described above.

DNA synthesis studies. Proliferating cells were identified by visualizing DNA synthesis in SFACs cultured for 24 h in H medium supplemented with 10 µM bromodeoxyuridine (BrdU; Sigma). Labeled cells were cytospun onto glass slides, fixed in ice-cold methanol, incubated with anti-BrdU antibody (Roche), and identified using the avidin-biotin complex immunoperoxidase method (Vector Laboratories, Burlingame, CA).

Cell culture photography. All images were captured with a 35-mm film camera or an Olympus MagnaFire S99806 digital camera (Olympus, Melville, NY). Both cameras were mounted on an Olympus IMT2 inverted microscope.

Electron microscopy. SFACs were fixed with 2.5% glutaraldehyde in 0.1% sodium cacodylate buffer at pH 7.4. After fixation, they were washed in 0.1% sodium cacodylate buffer pH 7.4, postfixed in 1% osmium tetroxide, and processed for Epon 812 (Ted Pella, Redding, CA) embedding. Thin sections were stained with uranyl acetate and lead citrate and examined with a JEOL transmission electron microscope.

Immunofluorescence staining. Before staining, nonadherent, undifferentiated SFACs were cytospun onto glass slides and fixed with 10% buffered formalin (Fisher Scientific, Middletown, VA). Adherent cells were fixed directly in 10% buffered formalin. Nonspecific antibody binding was blocked by incubating cells in PBS-0.1% Tween 20 containing 5% goat serum. Primary antibodies directed against CD34, Tie-2, PDGF receptor (PDGFR)-{alpha}, PDGFR-{beta} (Santa Cruz Biotechnology, Santa Cruz, CA), {alpha}-SMA, calponin, desmin (NeoMarkers, Fremont, CA), NG2 (Chemicon International), and CD31 were used. Parallel cultures were incubated with negative control IgG. Secondary antibodies were conjugated with Alexa Fluor 488 and Alexa Fluor 568 (Molecular Probes, Eugene, OR). Cells were mounted in Gelvatol (Monsanto, St. Louis, MO), and images were obtained with either a Leica TCS-SP laser-scanning confocal microscope or an Olympus BX41 fluorescence microscope equipped with an Optronics MicroFire SE digital camera.

RT-PCR. Total RNA was extracted from cells with the RNeasy Micro kit (Qiagen, Germany) and examined for both quality and quantity with a BioAnalyzer 2100 (Agilent, Palo Alto, CA). Random primed reverse transcription (RT) was performed with 200 ng of RNA and Superscript III reverse transcriptase (Invitrogen). Reactions lacking enzyme were performed in tandem for each RNA sample to act as negative controls.

For each sample, 1/20 RT reaction was used as a template for PCR. Reaction conditions were 10 mM Tris·HCl, 50 mM KCl, 0.1% Triton X-100, 250 µM 2-deoxynucleotide 5'-triphosphate, 1.5 mM (or 1 mM for CD34) MgCl2; 1 U of Taq DNA polymerase (Promega); and 0.5 µM each of forward and reverse primers (Table 1) in a 20-µl volume. PCR was performed at 95°C for 3 min, followed by 30 cycles of 94°C for 45 s, 56°C for 45 s, and 72°C for 40 s. PCR products were separated by performing electrophoresis in agarose gels containing ethidium bromide and visualized under UV light. All PCR products were sequenced to verify their identity.


View this table:
[in this window]
[in a new window]
 
Table 1. Oligonucleotide primers used to amplify genes of interest in SFACs

 

    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Rat aorta contains immature cells capable of anchorage-independent growth. CD31-negative cells isolated from the aorta of Fischer 344 male rats were cultured in a serum-free growth medium originally optimized for culturing human neural stem cells (42, 47). Some of these cells grew in suspension and formed clusters, which gradually organized as compact spheroids (Fig. 1A). The remaining cells attached to the culture dish and were discarded. Spheroids were then allowed to expand and proliferate in suspension. After 2–3 wk in culture, up to 300 spheroids ranging in size from 100 to 500 µm were obtained from a single aorta. Trypsin treatment of spheroids produced a single-cell suspension that retained the capacity to continuously grow and form new spheroids. We determined that these anchorage-independent cells were capable of forming SFACs. Proliferation of SFACs was relatively slow as indicated by DNA synthesis studies showing BrdU uptake in a small fraction of the cells during a 24-h period (Fig. 1B).



View larger version (141K):
[in this window]
[in a new window]
 
Fig. 1. Rat aorta-derived mesenchymal cells form spheroids in suspension culture. CD31-negative cells isolated from the rat aorta and cultured in serum-free medium optimized for the culture of stem cells grew in suspension forming spheroidal colonies (A). Bromodeoxyuridine (BrdU) labeling over a 24-h period demonstrated that some spheroid-forming aortic cells (SFACs) were synthesizing DNA (B). Electron microscopy shows that SFACs were viable throughout the spheroidal structures (C). Cells at the edge of the spheroids had short surface microvilli coated with a fine fibrillar meshwork (D, arrows). Cells within the spheroids exhibited an immature phenotype and were surrounded by a lacy extracellular matrix (E, arrows). Adjacent cells were focally connected by small adherent-type junctions (F, arrow). Scale bars: A, 250 µm; B, 50 µm; C, 20 µm; D, 1 µm; E and F, 2 µm.

 
SFACs have an undifferentiated ultrastructural morphology. When examined by electron microscopy, SFACs exhibited nuclei with abundant euchromatin and no evidence of specific cell differentiation. Cytoplasmic organelles included rough endoplasmic reticulum, free ribosomes, mitochondria, and Golgi complexes. SFACs appeared viable at the periphery as well as within the cores of the spheroids (Fig. 1C). Surface cells lining the spheroids were flattened and had short microvilli coated with a fine fibrillar meshwork (Fig. 1D). A similarly delicate extracellular matrix was identified in the intercellular spaces of the spheroids (Fig. 1E). Adjacent cells were connected by small adherent-type junctions (Fig. 1F). Isolated cells not incorporated into spheroids showed similar features, except for their nucleus-cytoplasm ratio, which was significantly higher because of the small amount of cytoplasm and cytoplasmic organelles (data not shown).

SFACs express a protein repertoire indicative of an immature mesenchymal phenotype. Protein expression by SFACs was studied using immunofluorescent staining followed by confocal microscopy. SFACs expressed CD34 and Tie-2 (Fig. 2, A and B) but were negative for {alpha}-SMA and CD31 (Fig. 2, C and D), indicating an immature cell phenotype. SFACs were also positive for NG2, nestin, PDGFR-{alpha}, and PDGFR-{beta} (Fig. 2, EH).



View larger version (60K):
[in this window]
[in a new window]
 
Fig. 2. SFACs have an immature phenotype on the basis of immunocytochemistry. SFACs expressed CD34 (A) and Tie-2 (B) but showed no immunoreactivity for markers of smooth muscle cell differentiation (C; {alpha}-smooth muscle actin, {alpha}-SMA) or endothelial cell differentiation (D; CD31). SFACs also expressed NG2 (E), nestin (F), PDGF receptor (PDGFR)-{alpha} (G), and PDGFR-{beta} (H). Scale bars, 40 µM.

 
bFGF is required for SFAC anchorage-independent growth. To evaluate the growth requirements of SFACs, we performed growth factor withdrawal experiments. These studies demonstrated that the anchorage-independent growth of SFACs and their capacity to form spheroids required bFGF, because cells cultured in medium lacking bFGF failed to grow in suspension but attached to the surface of the dish, forming colonies (Fig. 3). Conversely, EGF or LIF withdrawal had no effect on the growth pattern of these cells (data not shown).



View larger version (90K):
[in this window]
[in a new window]
 
Fig. 3. Basic fibroblast growth factor (bFGF) is required for SFAC anchorage-independent growth. A: SFACs grew in suspension and formed spheroids in bFGF-containing serum-free medium. B: when bFGF was omitted from the medium, the cells did not form spheroids but attached to the culture surface and grew in colonies. Scale bars, 50 µM.

 
Serum induces irreversible transformation of SFACs into anchorage-dependent cells. The growth pattern and immunocytochemical features indicated that SFACs were immature cells and represented a progenitor cell population. To investigate their differentiation potential, we performed a series of studies in which SFACs were exposed to endothelial or smooth muscle differentiation factors. Treatment with VEGF or ANG I, which have been shown to induce endothelial differentiation (16, 18), had no effect, as demonstrated by the persistent absence of CD31 and von Willebrand factor and the lack of any morphological changes in the spheroids (data not shown). Because serum has been shown to regulate smooth muscle cell differentiation (27, 29), we performed studies in which SFACs were transferred from serum-free medium (Fig. 4A) to medium containing 10% FBS. Serum-treated SFACs attached to the surface of the culture dish within 48 h, acquired a mesenchymal appearance (Fig. 4B), and then proliferated to confluence (Fig. 4C). Terminal differentiation of these cells was confirmed by the observation that serum-treated cells lost their capacity to form spheroids, proliferated, and were unable to revert to a spheroid-forming phenotype when returned to serum-free medium (Fig. 4D). When trypsin treated and incubated in serum-free medium, these cells reattached and grew more slowly than cells cultured in 10% serum but did not revert to the spheroid phenotype (Fig. 4E). Unlike SFACs, control cultures of RAECs and rat tail fibroblasts (56) failed to grow as anchorage-independent cells and were unable to form spheroids (data not shown). These findings suggested that SFACs are anchorage-independent progenitor cells that terminally differentiated into anchorage-dependent cells upon exposure to serum.



View larger version (78K):
[in this window]
[in a new window]
 
Fig. 4. SFACs differentiate in response to serum. Upon exposure to serum, SFACs lost their original phenotype (A) and transformed into mesenchymal type cells that attached to the culture surface within 2 days (B). These cells proliferated and formed a confluent monolayer by day 12 (C). Returning the cells to serum-free medium, either in the original culture (D) or after trypsin treatment and replating (E), failed to restore the original spheroid growth pattern. Scale bars, 100 µM.

 
Serum-induced differentiation of SFACs causes changes in protein and mRNA expression indicative of a mural cell phenotype. To evaluate changes in protein expression associated with serum-induced phenotypic modulation, SFACs were grown in 10% serum for 48 h to induce attachment and proliferation and were then evaluated using immunofluorescence staining. Serum-treated SFACs became CD34 negative (Fig. 5A) and {alpha}-SMA positive (Fig. 5B), suggesting that they had differentiated from progenitor cells to mural cells. This conclusion was corroborated by additional studies demonstrating expression in these cells of the mural cell markers calponin (Fig. 5C) and desmin (Fig. 5D). They also remained CD31 negative (Fig. 5E), confirming their inability to differentiate into endothelial cells. Serum-treated SFACs remained positive for Tie-2, NG2, nestin, PDGFR-{alpha}, and PDGFR-{beta} (Fig. 5, FJ).



View larger version (64K):
[in this window]
[in a new window]
 
Fig. 5. SFACs differentiate into a mural cell phenotype in response to serum. Serum-treated SFACs lost CD34 (A) and acquired {alpha}-SMA (B), calponin (C), and desmin (D) expression while remaining negative for CD31 (E) and positive for Tie-2 (F), NG2 (G), nestin (H), PDGFR-{alpha} (I), and PDGFR-{beta} (J). 4–6-diamidino-2-phenylindole-stained nuclei are shown in blue. Scale bars, 40 µM.

 
To corroborate these results, the expression of selected genes was examined with RT-PCR on SFACs grown with or without 10% serum. These studies confirmed that upon treatment with serum, SFACs switched from a CD34-positive and {alpha}-SMA-negative phenotype to a CD34-negative and {alpha}-SMA-positive phenotype (Fig. 6A). RT-PCR also showed that SFACs were negative for the endothelial cell markers CD31, endothelial nitric oxide synthase (eNOS), and fetal liver kinase (Flk)-1 before and after exposure to serum. As predicted by the immunofluorescence studies, SFACs expressed Tie-2, NG2, nestin, PDGFR-{alpha}, and PDGFR-{beta} and remained positive for these markers after serum treatment.



View larger version (57K):
[in this window]
[in a new window]
 
Fig. 6. RT-PCR confirmed the capacity of SFACs to differentiate into mural cells in response to serum. The effect of serum on SFAC differentiation was evaluated by performing RT-PCR on selected genes of interest. A and B: gene expression in SFACs maintained in serum-free medium (A) or in medium treated with serum (B). C: positive controls for endothelial genes. In AC, the presence of reverse transcriptase (RT) is indicated by + or – at tops of columns. SFACs were CD34 positive and {alpha}-SMA negative when maintained in serum-free medium but transformed into a CD34-negative and {alpha}-SMA-positive cell type upon exposure to serum. Tie-2, NG2, nestin, PDGFR-{alpha}, and PDGFR-{beta} were present before and after differentiation. CD31, eNOS, and fetal liver kinase (Flk)-1 mRNA were not expressed before or after differentiation.

 
PDGF-BB transform SFACs into pericyte-like cells in collagen gel. To evaluate the capacity of SFACs to interact with an extracellular matrix in response to growth factor stimulation, the cells were suspended within a 3D collagen gel and treated with PDGF-BB or VEGF or were left untreated. After 24 h, the control cells as well as the VEGF-treated cells had migrated minimally from the spheroids into the surrounding matrix (Fig. 7, A and B). In contrast, PDGF-BB-treated cells had actively invaded the collagen gel, which they colonized extensively in a few days (Fig. 7C). PDGF-BB-induced migration occurred within 1–2 h from exposure to the growth factor. Control and VEGF- treated cells had rounded shapes and relatively few cytoplasmic processes (Fig. 7, D and E). Conversely, PDGF-BB-treated cells exhibited a markedly dendritic, pericyte-like morphology, characterized by long and branching cell processes (Fig. 7F).



View larger version (102K):
[in this window]
[in a new window]
 
Fig. 7. PDGF-BB stimulates SFAC outgrowth and pericyte-like transformation as shown in photomicrographs of SFACs embedded in collagen gels and cultured for 24 h in serum-free medium without growth factors (A and D) or with the addition of either 50 ng/ml VEGF (B and E) or 50 ng/ml PDGF-BB (C and F). PDGF-BB markedly stimulated the outgrowth of SFACs into the collagen matrix (C). PDGF-BB-treated SFACs exhibited a pericyte-like, dendritic morphology (F) compared with untreated control (D) or VEGF-treated cells (E). Scale bars: AC, 100 µM; DF, 25 µM.

 
SFACs promote mural thickening of neovessels in angiogenic cultures of rat aorta. To evaluate the capacity of SFACs to interact with a growing vasculature, these cells were cocultured with angiogenic outgrowths of rat aorta in a collagen gel. Control cultures consisted of rat aorta alone. Aortic rings in this system generated neovessels composed of a continuous layer of endothelial cells and surrounding pericytes. As they contacted the proliferating vasculature, SFACs migrated toward and surrounded the neovessels, giving rise to a markedly thicker coating of pericytes than those in control cultures of aortic explants alone (Fig. 8). The multilayering of pericytes obtained in SFAC aortic ring cocultures was in striking contrast to the single discontinuous coating of pericytes of the control neovessels (Figs. 8, A and C). Many of the extra pericytes in the cocultures appeared to originate from the SFACs. Observations rendered using phase-contrast microscopy were confirmed by double staining the cultures for isolectin B4, an endothelial cell marker, and NG2, a SFAC and pericyte marker (Fig. 8, B, D, and E).



View larger version (87K):
[in this window]
[in a new window]
 
Fig. 8. SFACs become pericytes when cocultured with angiogenic outgrowths of rat aorta. Photomicrographs of living cultures (A and C) and confocal images (B, D, and E) of endothelial cells labeled in red with isolectin B4 and pericytes labeled in green with anti-NG2 antibody of neovessels from rat aortic rings cultured in collagen gels without (A and B) or with SFACs (CE). Control neovessels were composed of endothelial tubes coated with a single discontinuous layer of pericytes (A, arrows), whereas neovessels cocultured with SFACs had a significantly thicker mural coating with multiple layers of pericytes (C, arrows). Double-fluorescence labeling confirmed the presence of an increased number of NG2-positive pericytes around the isolectin-B4-positive endothelial tubes in aortic SFAC cocultures (D and E) compared with neovessels of control cultures (B). Close interaction of the tip of a branching neovessel with an implanted spheroid suggests that SFACs contributed to the formation of a periendothelial mural cell coating (E) . Scale bars: A and C, 50 µM; B, D, and E, 40 µM.

 
SFACs transform into pericytes in angiogenic cultures of isolated endothelial cells. To provide further evidence that SFACs have the capacity to become pericytes in an otherwise pericyte-free system, these cells were cocultured with a pure population of RAECs in a collagen gel under conditions of angiogenic stimulation. RAECs cultured without SFACs formed branching capillary tubes in response to VEGF and bFGF (Fig. 9A). In the cocultures, SFACs migrated toward the capillary tubes and attached to the external surface of the endothelium, becoming pericytes (Fig. 9B). This situation was in contrast to the RAEC-only cultures, which had naked capillary tubes without a pericyte coating (Fig. 9A). Immunofluorescent staining confirmed that SFAC-derived NG2-positive cells interacted closely with the isolectin B4-stained endothelial cell tubes in the collagen gel, forming a pericyte coating (Fig. 9, D and F). Control cultures with RAECs alone had no NG2-positive pericytes (Fig. 9, C and E).



View larger version (161K):
[in this window]
[in a new window]
 
Fig. 9. Photomicrographs of living cultures (A and B) and digital confocal images (CF) of capillary tubes formed by isolated rat aortic endothelial cells (RAEC) in three-dimensional collagen gel cultures without (A, C, E) or with (B, D, F) SFACs. Capillary tubes formed in the absence of SFACs were composed of endothelial cells only (A), whereas capillary tubes cocultured with SFACs comprised both endothelial cells and pericytes (B, arrows). Representative confocal images of RAEC-SFAC cocultures (D and F) confirmed the presence of NG2-positive pericytes (green) in close association with isolectin B4-positive endothelial cells (red) compared with control cultures of RAECs alone, which had no pericytes (C and E). Scale bars: A and B, 100 µM; CF, 20 µM.

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study demonstrates that the postnatal rat aorta contains an immature subpopulation of anchorage-independent mesenchymal cells with pericyte progenitor features. We termed them SFACs on the basis of their capacity to form spheroids in suspension culture. SFACs express CD34 and Tie-2 and are negative for CD31, eNOS, Flk-1, and {alpha}-SMA, a protein repertoire compatible with a primitive, undifferentiated phenotype. They are also positive for markers of early pericyte lineage (NG2, nestin, PDGFR-{alpha}, and PDGFR-{beta}). Upon treatment with serum, SFACs lose their capacity to grow in suspension and differentiate into a CD34-negative, {alpha}-SMA-positive mural cell phenotype while remaining negative for markers of endothelial cell differentiation. In addition, they express the mural cell markers calponin and desmin. SFACs respond to PDGF-BB in 3D collagen gel culture by migrating and forming dendritic processes and transform into pericytes when cocultured with angiogenic outgrowths of rat aortic rings or isolated endothelial cells.

CD34-positive cells capable of smooth muscle cell differentiation have been identified in postnatal peripheral blood (58), small intestine (53), and skeletal muscle (22). There is also evidence that the embryonal aorta contains an abundant population of CD34+/CD31– cells (1), of which only a fraction differentiate into mature endothelial cells. The remaining cells may represent the embryonal lineage of the CD34-positive/CD31-negative SFACs that were isolated in this study from the postnatal rat aorta.

Serum-treated SFACs became CD34-negative and gained expression of mural cell markers but not CD31, eNOS, or Flk-1. This finding indicates that the SFACs were able to differentiate into mural cells but not into endothelial cells. Although the mechanisms regulating SFAC differentiation remain to be elucidated, it is possible that serum-mediated SFAC transformation into a mural cell phenotype is linked to the serum response factor, a serum-induced transcription factor that has been implicated in smooth muscle and skeletal muscle differentiation (27, 29).

Several observations in our study indicate that rat aorta-derived SFACs are pericyte progenitor cells. First, these cells proliferated in suspension-forming spheroids, a growth pattern associated with stem and/or progenitor cells (42, 49). Second, the rate of SFAC proliferation was relatively slow as is typically observed in progenitor cell populations. Third, the ultrastructural morphology of SFACs showed no evidence of features characteristic of a differentiated phenotype. Fourth, SFACs expressed CD34, a protein characteristically associated with stem cells (45). Finally, when cocultured with aorta-derived neovessels, SFACs gave rise to a multilayered coating of pericytes, which was much more complex than the single discontinuous layer of pericytes of control microvessels. The pericyte progenitor nature of SFACs was confirmed with an in vitro assay system in which capillary tubes formed by isolated endothelial cells became surrounded by SFAC-derived pericytes.

SFACs expressed both PDGF receptors {alpha} and {beta} and responded to PDGF-BB, which can bind and activate both receptors (15). PDGF-BB has been shown to play a critical role in pericyte recruitment during angiogenesis (8, 17, 20, 28). PDGFR-{alpha} appears to be particularly important for smooth muscle cells of large vessels (50), whereas PDGFR-{beta} is essential for the differentiation of microvascular pericytes (21, 54). Interestingly, NG2, which is strongly expressed in SFACs, binds to PDGF-AA and functions as a requisite coreceptor for the action of this ligand on PDGFR-{alpha} (13, 14).

The capacity of SFACs to grow in suspension and their ability to form spheroids was strictly dependent on bFGF. When bFGF was omitted from the medium, SFACs attached to the culture surface and appeared to differentiate. These findings are consistent with the observation that bFGF is often required to maintain stem cells in culture (34). It is noteworthy that NG2 acts as an auxiliary bFGF receptor and potentiates the ability of bFGF to interact with and activate its tyrosine kinase receptors (13).

SFACs are positive for Tie-2, which was originally reported as an endothelial cell-specific receptor (51). However, there is evidence that Tie-2 also can be expressed in mural cell lineages (23, 43, 55) and by nonendothelial mesenchymal cells during angiogenesis in vivo (32).

Of particular interest is that the isolation and propagation of SFACs was made possible by the use of a serum-free medium originally introduced for the culture of neural stem cells (42, 47). Our finding that gene expression profiles are shared by neural cells and SFACs points to molecular similarities between these cell types. For example, NG2 and nestin, which are both expressed in SFACs, were originally described as neural markers but were later identified in immature mesenchymal cells and mural cells (2, 40, 41, 57). The observation that expression of genes associated with neural cells are also found in SFACs suggest that there are developmental overlaps in the differentiation of these distinct cell types. The finding that avian cranial neuroectoderm can generate pericytes when transplanted in quail chicks (26) supports this concept. Moreover, although most vascular mural cells are of mesodermal origin, the smooth muscle cells originate in the aortic arch, and proximal great vessels originate from the neural crest (7, 19, 31).

In addition to providing a model of mural cell differentiation, SFACs represent a valuable alternative to conventional pericyte cell strains and lines. SFACs are relatively easy to isolate and can be passaged and grown in culture for several months. In contrast, mature pericytes are difficult to isolate and fail to grow beyond a few passages unless immortalized (4, 5, 25).

In summary, our studies have indicated that the postnatal rat aorta contains a population of immature mesenchymal cells with progenitor cell features. The suspension culture method described in this article can be used to isolate, maintain, and propagate these cells in an undifferentiated state for several months. Serum, exposure to PDGF-BB, or coculture with endothelial cells induces transformation of these cells into pericytes. These cells lend themselves to studies of mural cell differentiation because of their characteristic immature phenotype. They also represent a useful source of mural cells for vascular bioengineering applications. Finally, because of their ability to survive in suspension and interact with angiogenic endothelial cells, these cells may prove to be a valuable tool for in vivo homing studies designed to target peripheral tissues undergoing physiological or pathological angiogenic responses.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grant HL-52585 (to R. F. Nicosia) and a grant from the Medical Research Service, Department of Veterans Affairs (to R. F. Nicosia).


    ACKNOWLEDGMENTS
 
We gratefully acknowledge Debbie Jones, Division of Pathology and Laboratory Medicine, Veterans Affairs Puget Sound Health Care System (Seattle, WA), for excellent technical assistance with the electron microscopy studies.


    FOOTNOTES
 

Address for reprint requests and other correspondence: R. F. Nicosia, Division of Pathology and Laboratory Medicine (S-113-Lab), Veterans Affairs Puget Sound Health Care System, 1660 South Columbian Way, Seattle, WA 98108 (e-mail: roberto.nicosia{at}med.va.gov)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Alessandri G, Girelli M, Taccagni G, Colombo A, Nicosia R, Caruso A, Baronio M, Pagano S, Cova L, and Parati E. Human vasculogenesis ex vivo: embryonal aorta as a tool for isolation of endothelial cell progenitors. Lab Invest 81: 875–885, 2001.[ISI][Medline]

2. Alliot F, Rutin J, Leenen PJM, and Pessac B. Pericytes and periendothelial cells of brain parenchyma vessels co-express aminopeptidase N, aminopeptidase A, and nestin. J Neurosci Res 58: 367–378, 1999.[CrossRef][ISI][Medline]

3. Asahara T, Masuda H, Takahashi T, Kalka C, Pastore C, Silver M, Kearne M, Magner M, and Isner JM. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res 85: 221–228, 1999.[Abstract/Free Full Text]

4. Asashima T, Iizasa H, Terasaki T, Hosoya K, Tetsuka K, Ueda M, Obinata M, and Nakashima E. Newly developed rat brain pericyte cell line, TR-PCT1, responds to transforming growth factor-{beta}1 and {beta}-glycerophosphate. Eur J Cell Biol 81: 145–152, 2002.[CrossRef][ISI][Medline]

5. Asashima T, Iizasa H, Terasaki T, and Nakashima E. Rat brain pericyte cell lines expressing {beta}2-adrenergic receptor, angiotensin II receptor type 1A, klotho, and CXCR4 mRNAs despite having endothelial cell markers. J Cell Physiol 197: 69–76, 2003.[CrossRef][ISI][Medline]

6. Carmeliet P and Jain RK. Angiogenesis in cancer and other diseases. Nature 407: 249–257, 2000.[CrossRef][ISI][Medline]

7. Chen S and Lechleider RJ. Transforming growth factor-{beta}-induced differentiation of smooth muscle from a neural crest stem cell line. Circ Res 94: 1195–1202, 2004.[Abstract/Free Full Text]

8. Crosby JR, Seifert RA, Soriano P, and Bowen-Pope DF. Chimaeric analysis reveals role of Pdgf receptors in all muscle lineages. Nat Genet 18: 385–388, 1998.[CrossRef][ISI][Medline]

9. Elsdale T and Bard J. Collagen substrata for studies on cell behavior. J Cell Biol 54: 626–637, 1972.[Abstract/Free Full Text]

10. Espinosa-Heidmann DG, Caicedo A, Hernandez EP, Csaky KG, and Cousins SW. Bone marrow-derived progenitor cells contribute to experimental choroidal neovascularization. Invest Ophthalmol Vis Sci 44: 4914–4919, 2003.[Abstract/Free Full Text]

11. Ferrara N. Vascular endothelial growth factor: molecular and biological aspects. Curr Top Microbiol Immunol 237: 1–30, 1999.[ISI][Medline]

12. Folkman J. Role of angiogenesis in tumor growth and metastasis. Semin Oncol 29: 15–18, 2002.[Medline]

13. Goretzki L, Burg MA, Grako KA, and Stallcup WB. High-affinity binding of basic fibroblast growth factor and platelet-derived growth factor-AA to the core protein of the NG2 proteoglycan. J Biol Chem 274: 16831–16837, 1999.[Abstract/Free Full Text]

14. Grako KA, Ochiya T, Barritt D, Nishiyama A, and Stallcup WB. PDGF {alpha}-receptor is unresponsive to PDGF-AA in aortic smooth muscle cells from the NG2 knockout mouse. J Cell Sci 112: 905–915, 1999.[Abstract/Free Full Text]

15. Hart CE, Forstrom JW, Kelly JD, Seifert RA, Smith RA, Ross R, Murray MJ, and Bowen-Pope DF. Two classes of PDGF receptor recognize different isoforms of PDGF. Science 240: 1529–1531, 1988.[ISI][Medline]

16. Hattori K, Dias S, Heissig B, Hackett NR, Lyden D, Tateno M, Hicklin DJ, Zhu Z, Witte L, Crystal RG, Moore MA, and Rafii S. Vascular endothelial growth factor and angiopoietin-1 stimulate postnatal hematopoiesis by recruitment of vasculogenic and hematopoietic stem cells. J Exp Med 193: 1005–1014, 2001.[Abstract/Free Full Text]

17. Hellström M, Kalén M, Lindahl P, Abramsson A, and Betsholtz C. Role of PDGF-B and PDGFR-{beta} in recruitment of vascular smooth muscle cells and pericytes during embryonic blood vessel formation in the mouse. Development 126: 3047–3055, 1999.[Abstract/Free Full Text]

18. Hildbrand P, Cirulli V, Prinsen RC, Smith KA, Torbett BE, Salomon DR, and Crisa L. The role of angiopoietins in the development of endothelial cells from cord blood CD34+ progenitors. Blood 104: 2010–2019, 2004.[Abstract/Free Full Text]

19. Hirschi KK and Majesky MW. Smooth muscle stem cells. Anat Rec A Discov Mol Cell Evol Biol 276: 22–33, 2004.[Medline]

20. Hirschi KK, Rohovsky SA, Beck LH, Smith SR, and D'Amore PA. Endothelial cells modulate the proliferation of mural cell precursors via platelet-derived growth factor-BB and heterotypic cell contact. Circ Res 84: 298–305, 1999.[Abstract/Free Full Text]

21. Hirschi KK, Rohovsky SA, and D'Amore PA. PDGF, TGF-{beta}, and heterotypic cell-cell interactions mediate endothelial cell-induced recruitment of 10T1/2 cells and their differentiation to a smooth muscle fate. J Cell Biol 141: 805–814, 1998.[Abstract/Free Full Text]

22. Hwang JH, Yuk SH, Lee JH, Lyoo WS, Ghil SH, Lee SS, Khang IG, Paik SY, and Lee JY. Isolation of muscle derived stem cells from rat and its smooth muscle differentiation. Mol Cells 17: 57–61, 2004.[ISI][Medline]

23. Iurlaro M, Scatena M, Zhu WH, Fogel E, Wieting SL, and Nicosia RF. Rat aorta-derived mural precursor cells express the Tie2 receptor and respond directly to stimulation by angiopoietins. J Cell Sci 116: 3635–3643, 2003.[Abstract/Free Full Text]

24. Kawasaki S, Mori M, and Awai M. Capillary growth of rat aortic segments cultured in collagen gel without serum. Acta Pathol Jpn 39: 712–718, 1989.[Medline]

25. Kondo T, Hosoya K, Hori S, Tomi M, Ohtsuki S, Takanaga H, Nakashima E, Iizasa H, Asashima T, Ueda M, Obinata M, and Terasaki T. Establishment of conditionally immortalized rat retinal pericyte cell lines (TR-rPCT) and their application in a co-culture system using retinal capillary endothelial cell line (TR-iBRB2). Cell Struct Funct 28: 145–153, 2003.[CrossRef][ISI][Medline]

26. Korn J, Christ B, and Kurz H. Neuroectodermal origin of brain pericytes and vascular smooth muscle cells. J Comp Neurol 442: 78–88, 2002.[CrossRef][ISI][Medline]

27. Landerholm TE, Dong XR, Lu J, Belaguli NS, Schwartz RJ, and Majesky MW. A role for serum response factor in coronary smooth muscle differentiation from proepicardial cells. Development 126: 2053–2062, 1999.[Abstract/Free Full Text]

28. Lindahl P, Johansson BR, Leveen P, and Betsholtz C. Pericyte loss and microaneurysm formation in PDGF-B-deficient mice. Science 277: 242–245, 1997.[Abstract/Free Full Text]

29. Mack CP, Thompson MM, Lawrenz-Smith S, and Owens GK. Smooth muscle {alpha}-actin CArG elements coordinate formation of a smooth muscle cell-selective, serum response factor-containing activation complex. Circ Res 86: 221–232, 2000.[Abstract/Free Full Text]

30. Majka SM, Jackson KA, Kienstra KA, Majesky MW, Goodell MA, and Hirschi KK. Distinct progenitor populations in skeletal muscle are bone marrow derived and exhibit different cell fates during vascular regeneration. J Clin Invest 111: 71–79, 2003.[Abstract/Free Full Text]

31. Mann KM, Ray JL, Moon ES, Sass KM, and Benson MR. Calcineurin initiates smooth muscle differentiation in neural crest stem cells. J Cell Biol 165: 483–491, 2004.[Abstract/Free Full Text]

32. Metheny-Barlow LJ, Tian S, Hayes AJ, and Li LY. Direct chemotactic action of angiopoietin-1 on mesenchymal cells in the presence of VEGF. Microvasc Res 68: 221–230, 2004.[CrossRef][ISI][Medline]

33. Montesano R, Orci L, and Vassalli P. In vitro rapid organization of endothelial cells into capillary-like networks is promoted by collagen matrices. J Cell Biol 97: 1648–1652, 1983.[Abstract]

34. Murphy M, Reid K, Ford M, Furness JB, and Bartlett PF. FGF2 regulates proliferation of neural crest cells, with subsequent neuronal differentiation regulated by LIF or related factors. Development 120: 3519–3528, 1994.[Abstract/Free Full Text]

35. Nicosia RF and Ottinetti A. Growth of microvessels in serum-free matrix culture of rat aorta: A quantitative assay of angiogenesis in vitro. Lab Invest 63: 115–122, 1990.[ISI][Medline]

36. Nicosia RF, Tchao R, and Leighton J. Histotypic angiogenesis in vitro: light microscopic, ultrastructural, and radioautographic studies. In Vitro 18: 538–549, 1982.[ISI][Medline]

37. Nicosia RF, Villaschi S, and Smith M. Isolation and characterization of vasoformative endothelial cells from the rat aorta. In Vitro Cell Dev Biol Anim 30A: 394–399, 1994.

38. Nicosia RF, Zhu WH, Fogel E, Howson KM, and Aplin AC. A new ex vivo model to study venous angiogenesis and arterio-venous anastomosis formation. J Vasc Res 42: 111–119, 2005.[CrossRef][ISI][Medline]

39. Nissanov J, Tuman RW, Gruver LM, and Fortunato JM. Automatic vessel segmentation and quantification of the rat aortic ring assay of angiogenesis. Lab Invest 73: 734–739, 1995.[ISI][Medline]

40. Ozerdem U, Grako KA, Dahlin-Huppe K, Monosov E, and Stallcup WB. NG2 proteoglycan is expressed exclusively by mural cells during vascular morphogenesis. Dev Dyn 222: 218–227, 2001.[CrossRef][ISI][Medline]

41. Ozerdem U, Monosov E, and Stallcup WB. NG2 proteoglycan expression by pericytes in pathological microvasculature. Microvasc Res 63: 129–134, 2002.[CrossRef][ISI][Medline]

42. Pagano SF, Impagnatiello F, Girelli M, Cova L, Grioni E, Onofri M, Cavallaro M, Etteri S, Vitello F, Giombini S, Solero CL, and Parati EA. Isolation and characterization of neural stem cells from the adult human olfactory bulb. Stem Cells 18: 295–300, 2000.[Abstract/Free Full Text]

43. Park YS, Kim NH, and Jo I. Hypoxia and vascular endothelial growth factor acutely up-regulate angiopoietin-1 and Tie2 mRNA in bovine retinal pericytes. Microvasc Res 65: 125–131, 2003.[CrossRef][ISI][Medline]

44. Rafii S, Meeus S, Dias S, Hattori K, Heissig B, Shmelkov S, Rafii D, and Lyden D. Contribution of marrow-derived progenitors to vascular and cardiac regeneration. Semin Cell Dev Biol 13: 61–67, 2002.[CrossRef][ISI][Medline]

45. Rafii S, Shapiro F, Rimarachin J, Nachman RL, Ferris B, Weksler B, Moore MA, and Asch AS. Isolation and characterization of human bone marrow microvascular endothelial cells: hematopoietic progenitor cell adhesion. Blood 84: 10–19, 1994.[Abstract/Free Full Text]

46. Rajantie I, Ilmonen M, Alminaite A, Ozerdem U, Alitalo K, and Salven P. Adult bone marrow-derived cells recruited during angiogenesis comprise precursors for periendothelial vascular mural cells. Blood 104: 2084–2086, 2004.[Abstract/Free Full Text]

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

48. Risau W. Mechanisms of angiogenesis. Nature 386: 671–674, 1997.[CrossRef][ISI][Medline]

49. Romero-Ramos M, Vourc'h P, Young HE, Lucas PA, Wu Y, Chivatakarn O, Zaman R, Dunkelman N, El-Kalay MA, and Chesselet MF. Neuronal differentiation of stem cells isolated from adult muscle. J Neurosci Res 69: 894–907, 2002.[CrossRef][ISI][Medline]

50. Schatteman GC, Motley ST, Effmann EL, and Bowen-Pope DF. Platelet-derived growth factor receptor {alpha} subunit deleted Patch mouse exhibits severe cardiovascular dysmorphogenesis. Teratology 51: 351–366, 1995.[ISI][Medline]

51. Schlaeger TM, Bartunkova S, Lawitts JA, Teichmann G, Risau W, Deutsch U, and Sato TN. Uniform vascular-endothelial-cell-specific gene expression in both embryonic and adult transgenic mice. Proc Natl Acad Sci USA 94: 3058–3063, 1997.[Abstract/Free Full Text]

52. Schor AM, Schor SL, and Allen TD. Effects of culture conditions on the proliferation, morphology and migration of bovine aortic endothelial cells. J Cell Sci 62: 267–285, 1983.[Abstract/Free Full Text]

53. Suárez-Rodríguez R and Belkind-Gerson J. Cultured nestin-positive cells from postnatal mouse small bowel differentiate ex vivo into neurons, glia, and smooth muscle. Stem Cells 22: 1373–1385, 2004.[Abstract/Free Full Text]

54. Tallquist MD, French WJ, and Soriano P. Additive effects of PDGF receptor {beta} signaling pathways in vascular smooth muscle cell development. PLoS Biol 1: e52, 2003.[Medline]

55. Tian S, Hayes AJ, Metheny-Barlow LJ, and Li LY. Stabilization of breast cancer xenograft tumour neovasculature by angiopoietin-1. Br J Cancer 86: 645–651, 2002.[CrossRef][ISI][Medline]

56. Villaschi S and Nicosia RF. Paracrine interactions between fibroblasts and endothelial cells in a serum-free coculture model: modulation of angiogenesis and collagen gel contraction. Lab Invest 71: 291–299, 1994.[ISI][Medline]

57. Vogel W, Grünebach F, Messam CA, Kanz L, Brugger W, and Bühring HJ. Heterogeneity among human bone marrow-derived mesenchymal stem cells and neural progenitor cells. Haematologica 88: 126–133, 2003.[ISI][Medline]

58. Yeh ETH, Zhang S, Wu HD, Körbling M, Willerson JT, and Estrov Z. Transdifferentiation of human peripheral blood CD34+-enriched cell population into cardiomyocytes, endothelial cells, and smooth muscle cells in vivo. Circulation 108: 2070–2073, 2003.[Abstract/Free Full Text]

59. Zhu WH and Nicosia RF. The thin prep rat aortic ring assay: a modified method for the characterization of angiogenesis in whole mounts. Angiogenesis 5: 81–86, 2002.[CrossRef][Medline]