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Keywords:

  • Mesenchymal stromal cells;
  • Human thoracic aorta;
  • Multiorgan donor;
  • Angiogenesis;
  • Allograft

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

The clinical use of endothelial progenitor cells is hampered by difficulties in obtaining an adequate number of functional progenitors. This study aimed to establish whether human thoracic aortas harvested from healthy multiorgan donors can be a valuable source of angiogenic progenitors. Immunohistochemical tissue studies showed that two distinct cell populations with putative stem cell capabilities, one composed of CD34+ cells and the other of c-kit+ cells, are present in between the media and adventitia of human thoracic aortas. Ki-67+ cells with high growth potential were located in an area corresponding to the site of CD34+ and c-kit+ cell residence. We thus isolated cells (0.5 ∼ 2.0 × 104 aortic progenitors per 25 cm2) which, upon culturing, coexpressed molecules of mesenchymal stromal cells (i.e., CD44+, CD90+, CD105+) and showed a transcript expression of stem cell markers (e.g., OCT4, c-kit, BCRP-1, Interleukin-6) and BMI-1. Cell expansion was adequate for use in a clinical setting. A subset of cultured cells acquired the phenotype of endothelial cells in the presence of vascular endothelial growth factor (e.g., increased expression of KDR and von Willebrand factor positivity), as documented by flow cytometry, immunofluorescence, electron microscopy, and reverse transcription-polymerase chain reaction assays. An in vitro angiogenesis test kit revealed that cells were able to form capillary-like structures within 6 hours of seeding. This study demonstrates that thoracic aortas from multiorgan donors yield mesenchymal stromal cells with the ability to differentiate in vitro into endothelial cells. These cells can be used for the creation of an allogenic bank of angiogenic progenitors, thus providing new options for restoring vascularization at ischemic sites.

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


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

There is some evidence that the arterial wall is much more dynamic than ever before believed. Studies have shown that postnatal angiogenesis may occur by recruitment of bone marrow resident cells with properties of embryonal angioblasts, that is, endothelial progenitor cells (EPCs), that are released to circulate in the blood in response to various stimuli [1, 2]. Once mobilized in the blood, EPCs are supposed to participate in physiological and pathological arterial wall remodeling during their lifetimes [3]. Although EPC homing to the sites of ischemia and contributing to the formation of new vessels have been demonstrated [4], others have obtained discordant results in this respect [5, [6], [7]8].

Hu et al. [9] recently found abundant vascular progenitor cells in the adventitia of ApoE-deficient mice; these progenitors contributed to experimental atherosclerosis and did not originate from the bone marrow. Moreover, human mature endothelial cells have been unexpectedly found to contain a subpopulation of EPCs allegedly organized in a completely hierarchical manner [10]. Thus, apart from the unquestionable link with the hematopoietic system, at least some progenitors seem to be located within the adult vessel wall.

Consistent with this view, constitutively resident vascular progenitor cells have been isolated from the thoracic and abdominal aortas of healthy adult mice [11] and from the internal thoracic arteries of humans [12]. The demonstration that resident vascular progenitors are present in the vessel wall might have a deep impact on several vascular biology fields, including EPC-based cell therapies for arterial insufficiency and tissue engineering.

For practical purposes, the therapeutic value of bone-marrow-derived EPCs, as well as vascular wall resident EPCs in regenerative medicine and tissue engineering, is being undermined by difficulties in the recruitment of an adequate number of healthy progenitors or a drop in the number and possibly functional properties of progenitors with age [13, 14]. These shortages can be overcome by using fresh arterial segments harvested from heart-beating multiorgan donors, which are collected at vascular tissue-banking facilities. This being the background, we here aimed to investigate whether thoracic aortas obtained from healthy adult multiorgan donors contain resident progenitors, which can be expanded and differentiated in vitro into endothelial cells suitable for restoring vascularization at ischemic sites.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

Tissue Studies

Fresh thoracic aortas and femoral arteries were harvested from 10 heart-beating multiorgan donors (six females; mean age 32 years); after procurement and decontamination, arterial samples were routinely processed for histological examination. Details of the procurement and sampling procedures are reported elsewhere [15].

The immunohistochemical staining here used was similar to that described previously [15]. Five μm-thick sections from formalin-fixed, paraffin embedded arteries were stained with the monoclonal antibodies (MoAbs) listed in the supplemental online material. To detect the antigen-antibody reaction, the streptavidin-biotin-peroxidase complex technique was used followed by diaminobenzidine tetrahydrochloride as a substrate solution (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com). Negative controls were performed by omitting the primary antibodies.

Cell Isolation, Culture, and Characterization

Approximately 5-cm-long fresh thoracic aortic segments were used for cell isolation. The arterial samples were longitudinally cut, providing an exposed surface area measuring approximately 25–30 cm2. The overall number of mononuclear cells recovered from each isolation varied from 1.5 × 106 to 3.5 × 106. After plating, the cells yielded by the primary culture ranged between 0.5–1 × 106. Additional details on cell culture are provided in the supplemental online material.

To detect surface antigens, aliquots of cells were stained by MoAbs as previously described [16]. Cells taken at passages 3–5 were washed twice with phosphate-buffered saline (PBS) containing 2% fetal bovine serum (FBS) and stained for 20 minutes at room temperature using the following MoAbs: CD133-phycoerythrin (PE) (Miltenyi Biotec, Bergisch Gladbach, Germany, http://www.miltenyibiotec.com), CD117-fluorescein isothiocyanate (FITC), CD34-PE, CD44-FITC, CD90(Thy-1.2)-phycoerythrin-cyanine 5 (PC5), CD105-PE, CD166-PE, CD45-allophycocyanin (APC), CD146-PE, human leukocyte antigen (HLA) class I-FITC, and HLA class II (HLA-DR)-FITC (all from Beckman Coulter, Fullerton, CA, http://www.beckmancoulter.com).

To study coexpression of CD105, CD44, and CD90(Thy-1.2), cells were simultaneously incubated with CD105-PE, CD44-FITC, and CD90(Thy-1.2)-PC5 MoAbs. Negative controls were run with appropriate conjugated irrelevant antibodies. Samples were analyzed using a Cytomics FC500 Flow Cytometer equipped with two lasers for data acquisition (Beckman Coulter). Results were analyzed using the CXP Software (Beckman Coulter).

For parallel immunofluorescence studies, the cells were plated at 1 × 103 per cm2 in collagen biocoated slide chambers (BD Biosciences, San Diego, http://www.bdbiosciences.com), cultured until near confluence, fixed in 2% paraformaldehyde, and labeled for 45 minutes at 37°C with MoAbs against CD44 (BD Pharmingen, San Diego, http://www.bdbiosciences.com/index_us.shtml), CD90 (BD Pharmingen), CD105 (BD Pharmingen), CD166 (BD Pharmingen), anti-human smooth muscle actin (ASMA) (Sigma), and CD45 (DakoCytomation, Glostrup, Denmark, http://www.dakocytomation.com). For intracytoplasmic ASMA staining, 0.01% Triton X-100 was added during fixation. The samples were first incubated with ASMA for 60 minutes at room temperature and then again incubated for 60 minutes with FITC-conjugated polyclonal rabbit anti-mouse immunoglobulins (DakoCytomation). Samples were observed under a fluorescence microscope. Details are provided in the supplemental online material. As already described [17], cells were also collected in Eppendorf tubes immediately after detachment; pellets were processed for electron microscopy as described in the supplemental online material.

Total RNA was extracted from cells using TRIzol Reagent according to the manufacturer's instructions (Invitrogen, Carlsbad, CA, http://www.invitrogen.com). Reverse transcription reactions were performed in a 20-μl volume with 2 μg of total RNA using M-MLV Reverse Transcriptase (Invitrogen) following the manufacturer's protocol. Oligo(dT)12–18 primers (Invitrogen) were used for first strand synthesis (details are provided in the supplemental online material). The expression of the following mRNAs was investigated: OCT4, BMI-1, BCRP-1, Interleukin-6 (IL6), c-kit, CD133, KDR, and β2 microglobulin (control).

Angiogenic Differentiation

An in vitro angiogenesis assay was performed according to Oswald et al. [18]. Cells taken at passage 5 were cultured until near confluence for 7 days in Dulbecco's modified Eagle's medium (DMEM) plus 2% FBS with and without 50 ng/ml vascular endothelial growth factor (VEGF) (Sigma) as well as in DMEM plus 10% FBS (negative control).

Matrigel (BD Biosciences) was prepared following the manufacturer's instructions. We dispensed 5 × 103 cells cultured in the conditions specified above into each well; plates were then incubated at 37°C 5% CO2. Human umbilical vein endothelial cells (HUVEC) were used as a positive control. The formation of capillary-like structures was observed in a CKX41 Olympus (Tokyo, http://www.olympus-global.com) inverted microscope after 2, 4, 6, and 20 hours. Experiments were performed in triplicate.

To visualize features of endothelial differentiation, 20-hour matrigel assays were fixed in situ with buffered 2% paraformaldehyde for 1 hour at 4°C. Fixation, postfixation, dehydration, and embedding steps were performed dispensing the appropriate solution into each well. Once the resin polymerized, the blocks were removed from each well and carefully cut until the structures previously observed by inverted microscope were reached. The ultrathin sections were counterstained and analyzed by an FEI Tecnai 12 transmission electron microscope.

In parallel experiments, the cells were analyzed at flow cytometry using the following anti-human MoAbs: KDR-APC (R&D Systems Inc., Minneapolis, http://www.rndsystems.com) to detect surface expression of VEGF receptor 2 and von Willebrand factor (vWF) (DakoCytomation) to detect the cytoplasmic expression of vWF. In this latter case, to demonstrate whether VEGF could prompt mesenchymal stromal cells (MSCs) to differentiate into endothelium, cells were permeabilized with the IntraPrep Kit (Beckman Coulter) and then incubated with vWF MoAb for 1 hour at room temperature. After two washes with PBS, cells were incubated with FITC anti-mouse IgG (Beckman Coulter) for 30 minutes at room temperature. Samples were then washed twice over and incubated for 20 minutes with normal mouse Ig (Sigma) to saturate free anti-mouse IgG sites. CD105 PE was added and incubated for 20 minutes and washed twice. Analyses were performed as described above.

To visualize the expression of vWF at immunofluorescence, the cells, conditioned as described above, were seeded (5,000 cells per cm2) on glass coverslips placed in six-well plates. After fixation, samples were stained with a MoAb against vWF (1:50; DakoCytomation). Blocking, mounting, nucleus counterstaining, and observations were performed as described above. The mean percentage of vWF+ cells was calculated by counting positive cells on digital images taken from 10 randomly selected fields at a magnification of ×40. At least 100 4,6-diamidino-2-phenylindole+ cells were analyzed each sample. To verify whether VEGF treatment could modify the expression of KDR and CD133 mRNAs, cells cultured until near confluence in the presence of VEGF were processed for reverse transcription-polymerase chain reaction (RT-PCR) analysis as specified above.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

The Thoracic Aorta of Multiorgan Donors Shows Distinct CD34+ and C-kit+ Cell Populations Between the Media and Adventitia Layers

Unlike femoral arteries, which show a thin layer of CD34+ capillary-like structures at the border between the media and adventitia, immunohistochemical studies of human thoracic aortas showed a more complex network of CD34+ cells, which also extended into the adjacent media and adventitia layers. The CD34+ cells were arranged both as single elements and capillary structures with evident lumina. vWF immunostaining performed on adjacent tissue sections only stained some of the larger vascular channels. CD34+ vascular channels were also seen in the media, which adjacent tissue sections showed to be vWF. CD45+ inflammatory cells were seen in association with the CD34+ cell network. A few c-kit+ round stromal cells were observed in the adventitia close to the richly vascularized border. Ki-67, a marker of cell proliferation, was used to label cycling endothelial and stromal cells. Representative immunohistochemical results are shown in Figure 1A–1I.

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Figure Figure 1.. Immunohistochemical characterization of cell populations located in between the media and adventitia arterial wall layers. (A): Femoral artery; (B–I): Thoracic aorta. Scale bars are 50 μm.

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After Culturing, Cells from Thoracic Aortas of Multiorgan Donors Show the Characteristics of Mesenchymal Stromal Cells

Cell isolation was carried out successfully in 3 of the 10 cases. In the remaining 7 cases, cell cultures were compromised by fast growth of contaminant bacteria and fungi and were therefore discarded. Common causes of biological contamination (e.g., media, solutions, and incubator) were ruled out. On the other hand, microbiological assays performed on the solution used for delivering arterial grafts to the Cardiovascular Tissue Bank facility revealed the presence of the bacterial species and yeasts listed in supplemental online Table 1.

In the successful cases, cell colonies began to appear approximately 3–4 days after initial plating. Considering that the doubling time of isolated cells was 36 hours (experiments not shown), the number of resident mesenchymal stromal cells present in the primary isolate from a 25 cm2 sample of aorta can be estimated as 0.5 ∼ 2.0 × 104 cells. Virological assays for cytomegolovirus and Epstein-Barr virus as well as cytogenetic analyses proved negative (results not shown).

At the beginning, cell growth consisted of cells with two distinct morphologies: one having a fibroblastoid appearance and the other one with a more rounded appearance (Fig. 2A). After passaging, these rounded cells rapidly disappeared from culture. The elongated cells continued to proliferate even after numerous passages (Fig. 2B), and their appearance was morphologically indistinguishable from that of MSCs isolated from the human bone marrow.

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Figure Figure 2.. Morphology and flow cytometry characterization of mesenchymal stromal cells (MSCs). Morphology of MSCs at initial plating (A) and at passage 3 (B). Scale bars are 20 (A) and 50 (B) μm. At flow cytometry (C), thoracic aorta MSCs coexpress CD44, CD90, and CD105 surface molecules. Percentages and cytograms from a representative experiment. Abbreviation: PE, phycoerythrin.

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Triple flow cytometry immunostaining of the cultured cells taken at passages 3–5 revealed that these cells coexpressed molecules commonly found in MSCs, that is, CD44, CD90, and CD105. In particular, 85% of the cell populations were CD44+/CD90+; 95% of CD44+/CD90+ also expressed CD105. Considering the results altogether, more than 80% of the overall cell population simultaneously expressed CD44, CD90, and CD105 molecules (Fig. 2C, representative of three different experiments, and supplemental online Table 2). Single labeling experiments also showed that at least 80% of the cultured cells were CD166+; KDR, CD34, and CD133 were found expressed in negligible cell fractions (0.1%–5%), whereas c-kit+ cells were not detected at all. Cells were also negative for lineage markers, that is, CD45, which is expressed in mature hematopoietic cells, and CD146 and vWF, which are markers of differentiated endothelium. Immunologically, the cells expressed HLA class I antigens and were negative for HLA-DR.

Flow cytometry results were confirmed by immunofluorescent single staining performed on cells cultured on slide chambers (Fig. 3A–3F). In summary, the phenotype of the isolated cells was CD44+, CD90+, CD105+, CD166+, KDRlow, CD133low, CD34low, c-kit, CD45, CD146, and vWF.

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Figure Figure 3.. Immunofluorescence characterization of mesenchymal stromal cells cultured on slide chambers. Cells were stained with monoclonal antibodies directed against CD44 (A), CD90 (B), CD105 (C), CD166 (D), ASMA (E), and CD45 (F). Scale bars are 10 μm. Abbreviation: ASMA, anti-human smooth muscle actin.

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Ultrastructural analysis showed cells with large euchromatic nuclei, prominent nucleoli, and abundant cytoplasm containing numerous organelles, for example, mitochondria and dilated cisternae of rough endoplasmic reticulum, a few lipid droplets, and peripheral cytoplasmic collections of clear vesicles, vacuoles, and blisters (supplemental online Fig. I). RT-PCR showed that these cells constitutively expressed the embryonic stem cell marker OCT4, some molecules involved in stem cell critical regulatory pathways (i.e., c-kit, BCRP-1, IL6, BMI-1, and KDR), as well as, to a lower extent, hematopoietic stem cell transcripts (e.g., CD133, supplemental online Fig. II).

Thoracic Aorta Mesenchymal Stromal Cells Have In Vitro Angiogenic Abilities

After 2 and 4 hours of Matrigel incubation, no significant difference was observed among the experimental conditions here investigated. After 6 hours of incubation, the cells that had previously been conditioned with VEGF showed a more elongated shape and distinct, thin cytoplasmic projections sprouting from the cell periphery; after 20 hours, the cells appeared to be connected by thicker projections, thus forming an evident capillary-like network. These features were not observed when cells had been cultured in the absence of VEGF. As expected, HUVEC spontaneously aggregated in a capillary-like network when seeded on Matrigel (Fig. 4A–4D).

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Figure Figure 4.. Mesenchymal stromal cells aggregated in capillary-like structures when seeded on Matrigel. Cells were cultured for 7 days in the absence (A) or presence (B–D) of VEGF; (B) corresponds to 6 hours from Matrigel seeding; (C, D) to 20 hours from Matrigel seeding. Abbreviations: ctr, control; hrs, hours; VEGF, vascular endothelial growth factor.

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After 20 hours of cell incubation, in situ ultrastructural analysis of Matrigel matrices showed cell aggregates having features consistent with an early endothelial phenotype; these characteristics included the presence of loosely arranged vimentin-like intermediate filaments, collections of micropinocytotic vesicles, and caveolae, tight junctions; around the Golgi area, moderately electron-dense oval-shaped granules possibly representing immature Weibel-Palade bodies were also seen (supplemental online Fig. III); this feature was only observed when cells had been treated with VEGF (results not shown).

In parallel assays, indirect immunofluorescence flow cytometry revealed that KDR expression was significantly increased when cells had been conditioned with VEGF for 7 days (Fig. 5A). Moreover, single (Fig. 5B) and double (Fig. 5C) labeling experiments demonstrated that VEGF promoted the vWF cytoplasmic expression in more than 50% of the CD105+ cell population (Fig. 5C, representative of three different experiments); vWF expression was exclusively observed after cells had been treated with VEGF. The expression of vWF was confirmed by immunofluorescence staining of cells cultured on glass coverslips. In these experiments, 12% ± 3% of cells revealed intense cytoplasmic staining for vWF when cultured in the presence of VEGF; vWF expression was associated with a rounded morphology of the positive cells (Fig. 5D); control HUVEC showed an intense vWF dot-like staining (Fig. 5E). Although an appropriate experimental design would probably have proved these conclusions to be correct, RT-PCR showed CD133 downregulation and KDR increased transcript expression (Fig. 5F, 5G) in the cells treated with VEGF.

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Figure Figure 5.. VEGF promotes endothelial differentiation in mesenchymal stromal cells (MSCs). Flow cytometry analysis of KDR ([A], single labeling staining) and vWF ([B], single labeling staining) expression in MSCs before and after VEGF incubation; double labeling staining demonstrates cytoplasmic expression of vWF in CD105+ cells (C). (D): Immunofluorescence staining shows vWF expression (white arrow) in VEGF-treated MSCs; (E): vWF dot-like staining in control human umbilical vein endothelial cells. Reverse transcription-polymerase chain reaction possibly documenting CD133 downregulation (F) and KDR upregulation (G) as a consequence of VEGF exposure. Scale bars (C, D) are 10 μm. Abbreviations: ctr, control; FBS, fetal bovine serum; VEGF, vascular endothelial growth factor, vWF, von Willebrand factor.

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Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

Recent studies have indicated that the human arterial wall contains a distinct subtype of resident progenitor cell with vasculogenic properties [12]. These cells, which have been termed vascular wall-resident progenitor cells, extend the spectrum of the known endothelial progenitors, which until now included the bone-marrow-resident EPCs (CD133+, CD34+, and KDR+) and their circulating counterpart EPCs (CD34+, KDR+).

EPCs are promising candidates for clinical applications aimed at restoring blood supply in ischemic damaged tissues [1]. However, the number and function of progenitors significantly decreases with age [13, 14] and disease [4, 19]. Circulating EPCs are extremely low in number [4]. Recent studies indicate that EPCs may play a minimal role in neovascularization of tumors, vessel repair or normal vessel growth, and development [5, [6], [7]8]. All these considerations justify the search for adequate alternative sources of these cells for autologous and allogenic use.

In this paper, we focused on thoracic aortas from heart-beating, multiorgan donors, which are collected at specifically authorized tissue banking facilities [20]. This strategy was inspired by the possibility of collecting a fair number of fresh large-sized arteries commonly derived from healthy young donors with high standards of quality and safety. The implications for the clinical use of tissues and cells are evident.

Our tissue studies show that two distinct cell populations, one composed of CD34+ cells and the other of c-kit+ cells, are present in human thoracic aortas. It is worth noting that both surface markers are expressed in cells with stem cell capabilities.

Immunohistochemical tissue studies showed a well developed plexus of CD34+ small vessels at the border between the media and adventitia layers; some of the cells lining these vessels were also vWF+, thus showing they had been employed in mature endothelium. This area, also containing CD45+ inflammatory cells, topographically corresponds to the CD34+/CD31 cell layer recently observed in human inner thoracic arteries, which ring assay studies have demonstrated to be highly vasculogenic [12].

Extending previous observations, we here found that the human thoracic aorta contains a more organized CD34+ cell network. In addition to the vasculogenic area, CD34+ cells were also seen as elongated elements in adjacent portions of the media and adventitia. CD34+ vascular channels, which adjacent sections showed to be vWF, could also be found embedded within the media. A similar vascular organization composed of cells highly expressing human leukocyte antigen class I antigen [16] was also found in another study and may therefore suggest that distinct patterns of CD34+ cell aggregation exist in elastic and muscular arteries.

The fact that such CD34+ cells can be organized into evident vascular structures does not conflict with the possibility that they might be progenitors; in fact, vascular wall-resident EPCs are believed to be the most likely recruitable cells from formed vessels contributing to new vessel formation [6]. This view is in accordance with the concept of angiogenesis, which is defined as the sprouting of new blood vessels from the differentiated endothelium of pre-existing vessels [21].

A novel finding was the observation of cells expressing the stem cell surface marker c-kit. This cell subpopulation was found to be scattered round c-kit+ cells in the adventitia stroma close to the vasculogenic zone. C-kit, which is commonly associated with the more primordial cardiac stem cells [22], is also expressed by mast cells; however, ultrastructural examination of contiguous samples ruled out this possibility (results not shown).

An additional finding comes from results with ki-67 tissue immunostaining. Ki-67, which recognizes a nuclear protein expressed in the G1, S, M, and G2 phases of the cell cycle, exclusively stained individual endothelial cells and stromal cells located between the media and adventitia. Although extensively looked for, other cells composing the arterial wall proved negative. Ki-67 tissue immunostaining documents that cells with high growth potential are located in a restricted area of the aortic wall corresponding to the site of CD34+ and c-kit+ cell residence.

It remains to be ascertained whether these cells could have angiogenic potential and whether such a property attaches to the endothelial cell side or to a closely related, adjacent stem-like cell that could be a perivascular pericyte [23] or an equivalent of the mouse and chicken mesoangioblast [24]. At a later stage, we succeeded in isolating a cell population from the thoracic aortas of multiorgan donors, which shared many properties with mesenchymal stromal cells [25].

Propagation of cells from 3 out of 10 specimens was achieved (30%); unsuccessful cases were caused by fast growth of bacteria as well as fungi at initial plating. These cultures were discarded to avoid delivering erroneous experimental results. We identified the solution used for arterial graft delivery to the Cardiovascular Tissue Bank facility as the vehicle of contamination; it is therefore evident that currently used arterial graft disinfection protocols, that is, 72 hours in an antibiotic mixture (mefoxin 240 mg/ml, lincomycin 120 mg/ml, colimycin 100 mg/ml, vancomycin 50 mg/ml), are not adequate for establishing cultures of arterial wall progenitors. This area needs further improvement.

Under culture conditions, the cells had a fibroblast-like appearance; flow cytometry showed that more than 80% of the cells coexpressed molecules commonly found in MSCs such as CD44, CD90, and CD105. Although performed as a single staining, CD166 was found expressed in most cells. Again, ultrastructural investigation revealed a strict resemblance with MSCs. After culturing, these cells showed a multivacuolar appearance and a high synthetic competence, that is, prominence of rough endoplasmic reticulum, as we recently observed in MSCs derived from the human bone marrow [17].

It is noteworthy that, in addition to such analogies, RT-PCR analysis revealed that these cells expressed transcripts associated with stem cells; some of these molecules are indeed involved in stem cell specific functions, such as proliferation and differentiation of hematopoietic progenitor cells (c-kit), maintaining the undifferentiated state of embryonic stem cells (IL6 and BCRP-1), regulating self-renewal of hematopoietic and neural stem cells (BMI-1), and governing pluripotency and cell fate determination of embryonic stem cells (OCT4) [26]. The expression of the BCRP-1 molecule is also linked to the specific ability of stem cells to exclude dyes such as rhodamine and Hoechst. This property, which identifies a small subset of stem cells termed the “side population” (SP), is due to the expression of transporter proteins, such as BCRP-1, which was found to be expressed in our study.

The existence of MSCs in adult arteries is supported by a recent paper documenting the isolation of an SP of stem cells from adult aortas in healthy mice [11]. Interestingly, these cells were able to differentiate into endothelial and smooth muscle cells but failed to give rise to hematopoietic lineages.

The presence of adult vascular wall-resident MSCs has been postulated in humans, too [12]. The authors there proposed a hypothetical scheme of the so-called “vasculogenic zone,” sketching a complete hierarchy of resident stem cells niched in the arterial wall (Fig. 10 in [12]). Within that proposed network of progenitors, we believe that the cells we have isolated could correspond to MSCs having a higher differentiation potential than EPCs.

At present, we do not know whether the MSCs we have isolated are multipotent; this aspect of their biology is under investigation. However, it should be noted that “ectopic” tissues (e.g., cartilage, bone, and fat) can be seen especially in atherosclerotic arteries during routine histopathological observations [27], and MSCs isolated from the human saphena vein are able to differentiate in vitro into osteoblasts, chondrocytes, and adipocytes [28]. Moreover, calcifying vascular cells isolated from the bovine aortic media display multilineage potential in vitro [27]. If the multilineage differentiation capacity of these cells is documented, the arteries from multiorgan donors may serve as an important source of multipotent MSCs for clinical needs and tissue engineering.

Meanwhile, these MSCs have shown the ability to differentiate into endothelium in vitro. In accordance with Oswald et al. [18], we used a three-dimensional Matrigel semisolid matrix to assay angiogenesis. After 20 hours of culture, the MSCs were able to form capillary-like structures, but this ability was strictly dependent on VEGF induction; controls always proved negative. VEGF induction was accompanied by increased cell expression of KDR, as documented by flow cytometry and RT-PCR assays; more importantly, we found that more than 50% of CD105+ cells coexpressed vWF following VEGF induction, as indicated by double labeling experiments performed at flow cytometry. At immunofluorescence, vWF intensely stained the cytoplasm of cells with a rounded morphology (12% ± 3%). This pattern of staining—already described in EPCs derived from chord blood mononuclear cells [29] as well as from amniotic membrane-derived MSCs [30]—was interpreted as a feature of early endothelial cell differentiation. Likewise, electron microscopy of the 20-hour samples showed features consistent with a basic endothelial cell employment, that is, collections of micropinocytotic vesicles and caveolae, tight junctions, and immature Weibel-Palade bodies.

The possibility of obtaining angiogenic MSCs from the thoracic aortas of multiorgan donors paves the way for creation of an allogenic bank of vascular progenitors of aortic origin. As already indicated, the fundamental scarcity of EPCs in the hematopoietic system is their main limitation when it comes to clinical application. Ex vivo expansion of EPCs cultured from the peripheral blood of healthy human volunteers yields approximately 5.0 × 106 cells per 100 milliliters of blood, whereas the amount of autologous bone marrow blood aspirated for therapeutic neovascularization is reported to be approximately 500 milliliters per person [31]. These figures suggest that at least 30 ∼ 45 × 107 EPCs are required in a clinical setting. Thus, the volume of blood required to extract an adequate number of progenitors for transplantation represents a practical limitation for the use of same. Under our experimental conditions, it can be calculated that 0.5 ∼ 2.0 × 104 progenitors are originally present in the mononuclear fraction obtained from a 5-cm-long segment of human thoracic aorta; these cells expand quickly and, a week after initial plating, their expansion yields approximately 0.5–1 × 106 cells. Considering the rate of expansion from the initial number of cells yielded, ideally the number of cells that could be obtained from a few in vitro passages might be approximately 1 × 108 cells, a reasonable amount for a clinical therapeutic approach to angiogenic disorders.

In summary, our findings indicate that thoracic aortas from heart-beating multiorgan donors are highly suitable for obtaining MSCs with the ability to differentiate in vitro into endothelial cells. Even though their differentiating potential remains to be fully established, it is believed that their angiogenic ability is in itself a useful property for allogenic use. These cells can be expanded rapidly, providing numbers that are adequate for therapeutic neovascularization; again, being recovered from young and healthy donors, they can be cryostored in appropriate cell banking facilities for later use.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

This work was supported by a grant from Ministero dell'Istruzione dell'Università e della Ricerca (http://www.miur.it) 2005 (diabetes and lower limb critical ischemia; study on arterial wall damage and the role of endothelial progenitor cells in artery repair and neoangiogenesis) and by a Grant from the University of Bologna (RFO number 6208, 2005).

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosure of Potential Conflicts of Interest
  8. Acknowledgements
  9. References
  10. Supporting Information
FilenameFormatSizeDescription
Figure_SI_pasquinelli.pdf687KSupplemental Figure 1
Figure_SII_pasquinelli.pdf22KSupplemental Figure 2
supplement_pasquinelli_R1.pdf31KSupplemental Data

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