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,,1,
,,2,,3
* Department of Pathology,
Department of Dermatology,
Department of Immunobiology, and
Interdepartmental Program in Vascular Biology and Transplantation, Yale University School of Medicine, New Haven, Connecticut, USA
2Correspondence: Yale University School of Medicine, 295 Congress Ave., Boyer Center for Molecular Medicine Rm. 454, New Haven, CT 06510, USA. E-mail: jordan.pober{at}yale.edu
ABSTRACT
We seeded tissue engineered human skin substitutes with endothelial cells (EC) differentiated in vitro from progenitors from umbilical cord blood (CB-EC) or adult peripheral blood (AB-EC), comparing the results to previous work using cultured human umbilical vein EC (HUVEC) with or without Bcl-2 transduction. Vascularized skin substitutes were prepared by seeding Bcl-2-transduced or nontransduced HUVEC, CB-EC, or AB-EC on the deep surface of decellularized human dermis following keratinocyte coverage of the epidermal surface. These skin substitutes were transplanted onto C.B-17 SCID/beige mice receiving systemic rapamycin or vehicle control and were analyzed 21 d later. CB-EC and Bcl-2-HUVEC formed more human EC-lined vessels than AB-EC or control HUVEC; CB-EC, Bcl-2-HUVEC, and AB-EC but not control HUVEC promoted ingrowth of mouse EC-lined vessels. Bcl-2 transduction increased the number of human and mouse EC-lined vessels in grafts seeded with HUVEC but not with CB-EC or AB-EC. Both CB-EC and AB-EC-induced microvessels became invested by smooth muscle cell-specific alpha-actin-positive mural cells, indicative of maturation. Rapamycin inhibited ingrowth of mouse EC-lined vessels but did not inhibit formation of human EC-lined vessels. We conclude that EC differentiated from circulating progenitors can be utilized to vascularize human skin substitutes even in the setting of compromised host angiogenesis/vasculogenesis.—Shepherd, B. R., Enis, D. R., Wang, F., Suarez, Y., Pober, J. S., Schechner, J. S. Vascularization and engraftment of a human skin substitute using circulating progenitor cell-derived endothelial cells.
Key Words: endothelial progenitor cell • umbilical cord blood • leukapheresis • angiogenesis • tissue engineering
TISSUE ENGINEERED HUMAN skin substitutes provide an alternative to skin autografts for treatment of nonhealing wounds that often develop in patients with vascular insufficiency (e.g., with venous stasis or diabetes or in the elderly). While skin substitutes such as Apligraf promote healing, they usually do not stably engraft and significant numbers of patients receiving bioengineered skin substitutes require additional interventions as a result of graft failure (1)
. Apligraf graft failure appears to be nonimmunological in that recipients do not become sensitized to human alloantigens expressed by graft cells. Instead, the primary reason for graft failure appears to be lack of adequate graft vascularization leading to hypoperfusion and ischemic injury (2
, 3)
. Usually, the survival of cells within the engrafted skin substitute is limited by diffusion of nutrients and oxygen from the underlying wound site, and this is generally inadequate for sustained survival (4)
. Blood vessel development within implanted avascular skin substitutes depends on host angiogenesis from microvessels in the wound bed and on vasculogenesis from circulating endothelial progenitor cells (5
6
7
8)
. Under optimal conditions, this process of graft vascularization requires at least 14 d to occur and is likely to be significantly delayed in patients with vascular insufficiency (9
, 10)
. In other words, the same conditions that predispose patients to the development of nonhealing skin ulcers reduce the efficiency of vascularization of skin substitutes and cause graft failure.
Novel strategies for vascularizing skin substitutes by incorporating endothelial cells (EC) have recently demonstrated encouraging preclinical and clinical results (11
12
13
14
15
16
17)
. We have reported a model of augmented vascularization of human skin substitutes formed from decellularized dermis and human neonatal foreskin keratinocytes induced by seeding the underside of the grafts with human umbilical vein EC (HUVEC) prior to orthotopic transplantation to the backs of immunodeficient (C.B-17 SCID/Bg) mice (18)
. Both formation of human EC-lined vessels within the engrafted construct and inosculation with mouse microvessels were demonstrated. The concentration of vascularization and the frequency of successful engraftment were increased further when the seeded HUVEC had been retrovirally transduced to constitutively express the antiapoptotic protein Bcl-2, a manipulation that increases the capacity of HUVEC to form mature vessels (19
, 20)
.
Although human EC improve the vascularization of skin substitutes, they potentially introduce a new complication, namely immunological rejection. Unlike keratinocytes or fibroblasts, human EC can initiate an immune response by resting allogeneic T-cells (21
, 22)
. Bcl-2 transduction makes HUVEC resistant to some (but not all) forms of immune-mediated injury (23)
. However, Bcl-2 transduced HUVEC may be more potent stimulators of alloimmunity than untransduced EC due to their increased survival and persistence in vivo (24)
. An alternative strategy to avoid an allogeneic response is to use EC autologous to the recipient to pre-seed the vascular component of the graft. It is generally impractical to derive such cells from the recipient’s own skin, especially in patients for whom skin harvesting may well create new nonhealing wounds. A potential approach to this obstacle is to derive EC from circulating endothelial progenitor cells (EPC). These cells, first described by Asahara et al. (25)
, have since been shown to actively participate in neovascularization and vascular adaptation in the adult (26
27
28)
. EPC may be isolated from either umbilical cord blood, adult bone marrow, or adult peripheral blood. The phenotype of these cells is not completely described, but the cells are enriched within populations that express CD34, CD133, and vascular endothelial growth factor (VEGF) receptor 2 (VEGFR2) (29
, 30)
. When cultured in the presence of VEGF, EPC lose CD133 expression, differentiate into cells that resemble other cultured EC types, and expand in number (31)
. True EPC, and their differentiated progeny, may be distinguished from monocytes (which may acquire EC markers when exposed to VEGF) by their much greater growth potential and their lack of leukocyte markers such as CD45 and CD14 (32
33
34)
.
In the present study, we have used our model of vascularized human skin substitutes to compare differentiated EC derived from culture of cord blood (CB) or adult blood (AB) EPC (designated CB-EC and AB-EC, respectively), with HUVEC. We have also examined the effect of exogenous Bcl-2 expression on CB-EC and AB-EC. Finally, we have compared the extent of vascularization produced by these various EC populations in a newly developed model of impaired host-angiogenesis/vasculogenesis produced by administration of rapamycin, an antiproliferative, immunosuppressive agent that inhibits angiogenesis in the mouse graft recipient. We conclude that AB-EC can be used to vascularize skin substitutes, although they are not as efficient as CB-EC, and that the beneficial effects of Bcl-2 on the potential of HUVEC to vascularize grafts are not observed in transduced populations of AB-EC or CB-EC. Importantly, seeded human EC still promote skin substitute vascularization in a setting of impaired recipient angiogenesis.
MATERIALS AND METHODS
Cell culture
All human cell populations were obtained using protocols approved by the Yale Human Investigation Committee. Keratinocytes were isolated from discarded neonatal human foreskins by dispase 0.025 g/mL in PBS (Roche Diagnostics, IN, USA) digestion, mechanical separation of the epidermis from the dermis, and further digestion of the isolated epidermis with trypsin-EDTA 0.05% (Life Technologies, Inc., Grand Island, NY, USA) for 3–5 min at room temperature (RT). The keratinocytes were then propagated in culture for one or two passages in KGM-2 media (Cambrex, Walkersville, MD, USA) prior to seeding of grafts. HUVEC were released from umbilical veins by collagenase digestion and serially cultured on 0.1% gelatin-coated flasks in M199/20% FBS supplemented with L-glutamine, penicillin/streptomycin (Life Technologies, Inc.), and endothelial cell growth supplement (ECGS, Calbiochem, La Jolla, CA, USA) with heparin from porcine intestine as described (35)
. EC were differentiated from EPC from umbilical cord blood or from adult peripheral blood. Collection of cord blood was performed immediately following elective cesarean section, and the blood was anticoagulated with heparin. Mononuclear cells were enriched by density centrifugation using Lymphocyte Separation Medium (MP Biomedicals, Aurora, OH, USA) according to manufacturer’s instructions. Peripheral blood mononuclear cells were collected from healthy volunteer adult donors by leukapheresis and enriched by density gradient centrifugation using Lymphocyte Separation Media. In the case of adult blood, CD34+ cells were further enriched by positive immuno-selection with antibody (Ab) and magnetic beads (MACSeparation Kit, Miltenyi, Auburn, CA, USA) according the manufacturer’s instructions. Total mononuclear cells from cord blood or CD34-expressing enriched mononuclear cells from adult peripheral blood were then plated onto gelatin-human plasma-fibronectin (0.1%, J.T. Baker, Phillipsburg, NJ, USA, and 20 µg/ml, isolated from outdated human plasma, respectively) coated tissue culture plastic and were cultured in EGM-2 media (Cambrex) supplemented with an additional 10 ng/ml VEGF (National Institutes of Health, Bethesda, MD, USA). Nonadherent cells were gently removed by washing after four days. Colonies of proliferating, differentiated cells were typically noted on days 7–10 (CB) or days 21–35 (AB), at which time the media was changed to EGM-2/15% FBS (FBS, Life Technologies) with no additional cytokine. Cultures were serially propagated on gelatin-coated flasks in EGM-2/15% FBS.
Characterization of EC
For analysis by immunofluorescence microscopy, cells were grown on glass coverslips coated with poly-L-lysine (Sigma, St. Louis, MO, USA) and 0.1% gelatin. Cells were rinsed once in PBS, fixed in 3.7% paraformaldehyde for 15 min at RT, and rinsed twice in PBS. The fixed monolayers were permeabilized in 0.1% Triton and then washed three times in PBS. Nonspecific binding sites were blocked by incubation with PBS containing 1% donkey serum (JacksonImmuno, West Grove, PA, USA) for 30 min at RT. Next, cells were incubated with the primary antibodies [antivon Willebrand Factor (vWF), 1:500, Dako, Carpinteria, CA, USA, for anti-VE-cadherin 1:100, Santa Cruz Biotechnology, Santa Cruz, CA, USA] diluted in PBS overnight at 4°C. After four washes in PBS, cells were incubated with secondary antibodies (for vWF, 1:500 goat anti-rabbit 568, Molecular Probes, Carlsbad, CA, USA; for VE-cadherin 1:500 goat anti- mouse AlexaFluor 488, Molecular Probes) for 45 min at RT, washed, and mounted with VectaShield (Vector Laboratories, Burlingame, CA, USA) with 4',6'-diam idino-2-phenylidole. Cells were imaged with a Zeiss Axiovert 2000M fluorescence microscope (Carl Zeiss, Inc., Thornwood, NY, USA).
Alternatively, cells were analyzed for surface expression of specific markers by fluorescence flow cytometry. Where indicated, cultured EC were exposed to TNF-
(10 ng/ml, R&D Systems, Minneapolis, MN, USA) for the times specified in the text prior to harvest. Confluent monolayers were washed twice in HBSS (HBSS, Life Technologies) and then incubated with trypsin-EDTA for 30–60 s. Enzyme activity was then quenched with 20% FBS/M199, and suspended cells were collected and washed in 10 ml, cold 1% BSA (BSA, Sigma)/PBS, centrifuged and washed again in cold 1% BSA/PBS. Cells were then incubated with 2 µg/ml primary Ab, or isotype control was directly conjugated to fluorescein isothiocyanate or Phycoerythrin for 45 min. Immunostained cells were then washed twice with cold PBS and analyzed on a FACSort using CellQuest software (BD Biosciences, San Jose, CA, USA). Specific antibodies used in these analyses were reactive with human CD18, CD31, CD34, CD45, CD133, CD62E, and VEGFR2 (CD18, CD31, CD45, and CD62E, 2 µg/ml, Beckman Coulter Immunotech, Miami, FL, USA; CD34 and CD133 2 µg/ml, Miltenyi; VEGFR2, 2 µg/ml, R&D Systems, Minneapolis, MN, USA). Alternatively, cells were stained with FITC-conjugated Ulex europeus agglutinin (Uea-1) (Sigma), which reacts with the blood group ABH expressed on human EC.
Transduction of EC
Stable transduction of HUVEC, CB-EC, or AB-EC with a caspase-resistant form of Bcl-2 was achieved as described previously for HUVEC (23)
. Briefly, subconfluent cultures of EC were infected daily for a total of four times with a supernatant containing the packaging virus D34A, Bcl-2 in the pSG5 expression vector, and Polybrene (Life Technologies, Inc.). The efficiency of transduction was confirmed by flow cytometry of permeabilized cells with anti Bcl-2 Ab, as described (23)
.
Preparation of engineered skin substitutes
A layer, 0.5 mM thick, of the epidermal-coated surface of cadaver donor skin, obtained from the Yale Skin Bank (discarded specimens), was prepared with a dermatome and then cut into 
5 cm x 5 cm pieces using a scalpel, rinsed in PBS (Life Technologies, Inc.) with antibiotics, subjected to three rapid freeze-thaw cycles in liquid nitrogen, and then incubated in PBS with antibiotics at 37°C for 2 wk, after which the epidermis was gently removed. Dermal pieces were further incubated in PBS with antibiotics for a total of 30 d at 37°C, then stored at –20°C until use. At this point, no residual viable cells could be identified by histological examination of the decellularized tissue.
Thawed 1 cm (2)
pieces of decellularized dermis were placed in 3 µl of KGM-2. After the dermis was rehydrated for at least 1 h at 37°C, the KGM-2 was removed. Keratinocytes (3x105), suspended in a 30 µl droplet of KGM-2, were pipetted onto the center of the former epidermal surface of the dermis and allowed to settle for 3 h. The graft was then covered with KGM-2, and the medium was changed to fresh KGM-2 on the following day. Three days after seeding, a differentiation medium was used consisting of KGM-2, Dulbecco’s modified Eagle medium, Ham’s F12 (FBS) chelated FBS, cholera toxin (1x10–10 M, Calbiochem), and hydrocortisone (0.4 µg/ml; BD Biosciences) plus amphotericin and gentomycin, with a final calcium concentration of 1.2 mM. The differentiation medium was changed every other day for 6–10 d until a multilayered keratinocyte covering had developed. At this time, the underside of the graft was seeded with cultured EC. Cells, 8 x 105 per graft, were introduced to the reticular dermis in a 30 µl droplet of M199/20% FBS/ECGS with hydrocortisone (0.4 µg/mL, cholera toxin (1x10–10 M), epidermal growth factor (10 ng/mL; BD Biosciences), and amphotericyn and gentimycin. After 3 h, more medium was added. Twenty-four hours after the grafts were seeded with EC, they were transplanted onto immunodeficient mice as described below.
Transplantation
All animal procedures were performed using protocols approved by the Yale IACUC. Three animals were used for each group. These groups were skin substitutes seeded with HUVEC, CB-EC, or AB-EC, and skin substitutes seeded with Bcl-2-transduced HUVEC, CB-EC, or AB-EC. Each group (±Bcl-2 transduction) was evaluated following systemic rapamycin treatment or treatment with vehicle control. Additionally, grafts constructed without EC seeding but with KC seeding were engrafted on animals receiving rapamycin treatment or treatment with vehicle. Analysis of engrafted skin substitutes was performed at 21 d.
Graft sites on the backs of female C.B-17 SCID/beige 6 to 10 wk of age (Taconic, Tarrytown, NY, USA) were prepared by first removing all visible hair from the graft site with a depilatory agent (NairTM, Carter-Wallace, New York, NY, USA). A 1 cm (2)
piece of mouse skin was removed to the concentration of the underlying fascia and a size-matched synthetic skin graft was sutured into the defect. The grafts were then covered with bacitracin ointment, and a waterproof, sutured dressing consisting of two layers of 1.5 cm2 TelfaTM (Kendall, Mansfield, MA, USA), TegadermTM (3M, St. Paul, MN, USA), a foam bandage (Stop and Shop, Boston, MA, USA), and a circumferentially wrapped DuroporeTM tape (3M). On the day of surgery and each day thereafter, 0.5–1.0 µl of PBS with L-glutamine, antibiotics, and amphotericin (Life Technologies, Inc.) was injected into the Telfa. The bandages were kept intact for 2 wk, at which time they were removed. Where indicated, animals were injected with rapamycin (3 mg/kg) or vehicle every day beginning on post-transplantation day 5. The mice were euthanized at 21 d, and grafts were harvested for analysis.
Histological analysis
Approximately half of each graft was fixed in formalin and paraffin-embedded for staining with hematoxylin and eosin (H&E) or immunohistochemistry. The other portion was snap-frozen in OCT (BD Biosciences, Franklin Lakes, NJ, USA) and 5 µm cryostat sections were prepared for immunohistochemistry.
Formalin-fixed paraffin-embedded tissue sections (6 µm) or frozen sections (5 µm) were used for immunohistochemical analysis of harvested tissue. Primary antibodies reactive with mouse CD31, human CD31, Bcl-2, or smooth muscle cell-specific alpha-actin were used for enumeration and characterization vascular profiles (mouse CD31, 1:100; human CD31 1:100; Bcl-2 1:100, BD Pharmingen; smooth muscle cell-specific alpha-actin 1:100 Novocastra, Newcastle on Tyne, UK). A biotinylated secondary Ab (1:100 JacksonImmuno, West Grove, PA) was used to detect the primary Ab and an avidin binding complex, and a 3-amino-9-ethyl carbazole detection kit was used for color development (Vector Laboratories).
Vessel density within engrafted skin substitutes was quantified by manual counting of a mid-graft tissue sample by an observer (BRS) blinded to the experimental protocol. Vascular profiles were characterized by positive staining with human or mouse CD31, in structures with an identifiable vascular lumen greater than 5 µm. All vessels in 10 random, high-power fields (54x54 µm) were counted for each graft. Statistical analysis was performed with a two-way ANOVA with a Bonferroni posthoc test.
RESULTS
Characterization of CB-EC and AB-EC
Using the conditions described in Materials and Methods, we observed that EC could be routinely differentiated from circulating progenitor cells found in either umbilical cord blood or peripheral blood from healthy adult volunteer donors. Unfractionated cord blood mononuclear cells gave rise to colonies of differentiated EC within 7 to 10 d of culture. Leukapheresis and enrichment of CD34+ cells was required for generation of colonies of differentiated, proliferating EC from adult peripheral blood and colonies were apparent only after 21–35 d of culture (Fig. 1
A, B). Although we did not formally assess EPC frequency, the requirement for concentration by leukapheresis and CD34 immunoselection is consistent with the conclusion that such cells are far more abundant in cord blood than in the peripheral blood of healthy adults not exposed to a stem cell mobilizing agent. Differentiated ECs from both donor sources formed a cobblestone-like monolayer of EC at confluence (Fig. 1C, D
) and could be serially expanded in vitro through multiple passages without obvious change in phenotype. By immunofluorescence microscopy, cultured EC derived from either CB or AB progenitors displayed VE-cadherin concentrated at the cell junctions and vWF in cytoplasmic granules (Fig. 1E, F
). Fluorescence flow cytometric analysis demonstrated that EC differentiated from either CB or AB progenitors were CD34, CD31, and VEGFR2 positive and bound Uea-1, all markers of differentiated EC (Fig. 2
A–D). These cells were also negative for CD133, a marker of undifferentiated EPC, and for CD45, CD18, and CD14, markers of monocytes retained by this cell type when cultured in VEGF (Fig. 2E-H
). In response to cytokine stimulation with TNF-, transient expression of E-selectin (CD62E) was observed in EC from circulating progenitors (Fig. 3
). By all these criteria, the cells used in our studies to seed synthetic skin are fully differentiated EC. Where indicated, cultured HUVEC, CB-EC, and AB-EC were transduced to express Bcl-2. On permeabilization, Bcl-2 could be detected in over 95% of transduced but not control EC by flow cytometry (Fig. 4
).
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Incorporation of CB-EC and AB-EC into skin substitutes
We next compared the abilities of CB-EC or AB-EC with that of HUVEC to produce blood vessels when incorporated into human skin substitutes prior to transplantation. At 21 d post-transplantation, control synthetic skin grafts containing a reconstructed epidermis consisted of human keratinocytes, but without seeding with EC, contained only a limited vasculature, which was largely restricted to the graft edges, presumably formed by invasion of the graft by host angiogenic microvessels (Fig. 5
A). As expected, all blood vessels in human keratinocyte-only control grafts were lined by ECs expressing murine CD31 but not human CD31. In animals treated with rapamycin, this limited degree of vascular infiltration was further reduced, leaving a largely avascular skin graft. In many animals with skin substitutes not seeded with human EC, rapamycin administration led to central necrosis and graft sloughing. In other words, systemic treatment of mice with rapamycin appeared to result in a state of compromised angiogenesis/vasculogenesis that frequently caused graft failure.
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Histological evaluation of skin substitutes constructed with either CB-EC or AB-EC revealed the presence of a well-vascularized dermis throughout the graft, with vessels extending into the dermal papillae (Fig. 5B, C
). The vessel density was consistently greater in CB-EC seeded grafts than in AB-EC seeded grafts and was at least comparable with the vessel density seen with Bcl-2-transduced HUVEC. The vessels formed by CB-EC and AB-EC, like those formed by HUVEC, acquired a coating of mural cells expressing smooth muscle cell-specific alpha-actin, indicative of vessel maturation (Fig. 5F
). The extent of mural cell investment in grafts containing CB-EC or AB-EC was also apparently unaffected by Bcl-2 transduction.
Grafts that were seeded with HUVEC were found to be well healed and vascularized (data not shown). As noted previously, the number of vessels within the graft was increased when using HUVEC that were transduced with a retrovirus encoding Bcl-2 compared to untransduced HUVEC (Fig. 6
A). Both human and mouse CD31-expressing cells were found lining the vessels of the engrafted skin substitutes, although human CD31-positive vascular profiles were more abundant at the center of the graft. Grafts containing Bcl-2 expressing HUVEC contained many more mouse EC-lined vessels than did grafts formed from untransduced HUVEC (Fig. 6A, D
). In other words, Bcl-2-transduced HUVEC (but not control, untransduced HUVEC) appeared to stimulate mouse angiogenesis, perhaps by release of paracrine factors. In animals treated with rapamycin and receiving skin grafts seeded with Bcl-2-transduced HUVEC, the number of vessels was reduced compared with vehicle controls (Fig. 6D
). Immunostaining revealed that rapamycin produced a marked reduction in the number of mouse EC-lined vessels within the graft, but had little effect on the number of human EC-lined vessels present compared to vehicle-treated mice. Rapamycin did not significantly reduce the numbers of vessels in synthetic skin grafts seeded with untransduced HUVEC, almost all of which are human EC-lined. In other words, experiments with HUVEC suggest that rapamycin effectively inhibits mouse angiogenesis but spares vessel formation mediated by incorporated human EC.
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AB-EC appeared to form at least as many vessels as untransduced HUVEC (Fig. 6B
). Bcl-2 transduction did not have a significant effect on the number of vessels formed by CB-EC or AB-EC (Fig. 6B, C
). When compared as groups, CB-EC generated significantly more human CD31-lined vessels than either HUVEC or AB-EC (Two-way ANOVA with Bonferroni posthoc test, P<0.001). Both CB-EC and AB-EC induced ingrowth of mouse CD31-positive vessels, and this was effectively suppressed by systemic administration of rapamycin (Fig. 6E, F
). The ability of CB-EC and AB-EC to induce host angiogenesis appeared comparable with that of Bcl-2-transduced HUVEC and was much greater than that induced by untransduced HUVEC. Although Bcl-2 was readily detectable by immunostaining in vessels lined by Bcl-2-transduced HUVEC, CB-EC, or AB-EC we did not detect Bcl-2 expression in untransduced CB-EC or AB-EC-lined vessels (data not shown), suggesting some additional mechanism other than Bcl-2 expression is responsible for the elaboration of proangiogenic signals from these cells.
DISCUSSION
In the present study we report the utilization of EC differentiated in vitro from circulating progenitor cells found in both human umbilical cord blood and in peripheral blood of healthy adult donors for the vascularization of tissue engineered human skin substitutes. Both CB-EC and AB-EC were found to be capable of promoting blood vessel formation in vivo when seeded on a skin substitute formed from decellularized human dermis and human keratinocytes, and then subsequently transplanted onto immunodeficient mice. Moreover, both CB-EC and AB-EC, like Bcl-2-transduced HUVEC, were not only able to form human EC-lined vascular structures within the skin substitutes but were also able to induce mouse angiogenesis within the grafts. By contrast, control HUVEC that were not transduced with the antiapoptotic protein Bcl-2 stimulated little host angiogenesis within the engrafted skin substitutes. In contrast to its effect on HUVEC, transduction of CB-EC or AB-EC with Bcl-2 did not appear to have a pronounced effect on either the number of human EC-lined vessels or mouse EC-lined vessels found within the skin grafts. Interestingly, CB-EC appeared to generate a greater number of total vessels (both human and mouse) than AB-EC (nontransduced or Bcl-2 transduced) or nontransduced HUVEC, while generating an equivalent number of total vessels to Bcl-2-transduced HUVEC.
As a model of impaired host angiogenesis, the antiproliferative immunosuppressive agent rapamycin was administered systemically to the mice following orthotopic transplantation of human skin substitutes. Rapamycin, inhibits processes dependent on cellular proliferation, including angiogenesis associated with wound healing (36
, 37)
. Indeed, delayed wound healing is a major side effect of this drug (38)
. Rapamycin administration suppressed mouse angiogenesis in synthetic skin grafts containing CB-EC, AB-EC, or Bcl-2-transduced HUVEC that otherwise showed enhanced host angiogenesis. The presence of rapamycin did not, however, impair the formation of human EC-lined vessels within the engrafted synthetic skin constructs. This preservation of human vascular contribution was demonstrated in all EC types evaluated in the current study. It is unclear why the drug did not affect vessel development by transplanted EC, but a possible explanation is that vessel formation by implanted EC is less dependent on cell division. Regardless of the explanation, this result bodes well for the use of EC transplantation to support new blood vessel development in the setting of impaired angiogenesis/vasculogenesis.
The results of the current study demonstrate that use of autologous AB-EC for vascularization of tissue engineered synthetic skin substitutes is feasible. Because such cells can theoretically be derived from potential graft recipients, they would not be expected to trigger allogeneic rejection responses. AB-EC like those differentiated from CB, show characteristic markers of EC in culture (Figs. 1
and 2)
. It is technically possible that the cells we isolated are actually circulating EC rather than EPC, since we did not select with CD133; however, the delay in colony formation and the subsequent vigorous expansion in culture is much more consistent with a stem cell precursor (34)
. In either case, the cells we are studying can be routinely isolated from blood and thus have the potential to avoid the need to harvest skin or other tissues. However, seeding of skin substitutes with CB-EC appears to lead to a more robust human vessel development than seeding with either AB-EC or HUVEC. CB-EC may, in fact, be a superior cell source for specific tissue engineering applications such as tissue engineered skin substitutes (39)
. At the present time, CB-EC are likely to be available only from allogeneic sources. We, therefore, expect skin substitutes made from CB-EC to stimulate a rejection response, perhaps requiring further genetic manipulation of these cells prior to use. This hypothesis is currently under investigation.
We have previously reported that retroviral transduction of HUVEC with Bcl-2 leads to enhanced vascularization of skin substitutes and maturation of human vessels within those engrafted constructs (18)
. Employment of Bcl-2 transduced CB-EC or AB-EC for graft seeding and vascularization did not lead to enhanced human (or mouse) EC-lined microvessels, nor did it seem to promote microvessel maturation at the 21 d time point. It is formally possible that Bcl-2 did exert an effect at an earlier time point not examined in this study. Even though Bcl-2 did not improve vessel development, it still may play a role in protecting allogenic CB-EC (or AB-EC) from rejection (24)
, a hypothesis under investigation in our laboratory. However, it is also possible that such cells, because they persist, will trigger an even stronger rejection response targeting other cell types within the graft that are not protected by immune injury. This question will require further study.
In summary, we have shown that human EC, differentiated and expanded in culture from circulating CB- or AB-EPC, can be used to promote the vascularization of synthetic skin substitutes. New blood vessels arise from both organization of the implanted human EC and from a recipient angiogenic/vasculogenic response. Most significantly, the ability of implanted cells to form new blood vessels is not compromised by the suppression of host angiogenesis/vasculogenesis, supporting this as a new strategy for the treatment of patients with impaired wound healing.
ACKNOWLEDGMENTS
We thank Bruce Fichandler and the Yale Skin Bank for providing human skin. We also thank Louise Benson, Gwen Davis, and Lisa Gras for assistance with cell culture and animal care and Dr. Edward Snyder and the Yale Blood Bank for assistance with leukaphereses. This work was supported in part by grafts from the Roche Organ Transplant Research Foundation (140097148 to J.S.S. and J.S.P.), the NIH (RO1 HL551014 and P30 AR41942 to J.S.P. and T32 HL07950 to B.R.S.), and Centro Nacional de Investigaciones Cardiovasculares Carlos III (CNIC Program 3+3 to Y.S.).
FOOTNOTES
1 Current address: Southern Illinois University School of Medicine, Department of Internal Medicine, Springfield, Illinois, USA. 
Received for publication January 6, 2006. Accepted for publication March 20, 2006.
REFERENCES
This article has been cited by other articles:
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  K. K. Hirschi, D. A. Ingram, and M. C. Yoder Assessing Identity, Phenotype, and Fate of Endothelial Progenitor Cells Arterioscler. Thromb. Vasc. Biol., September 1, 2008; 28(9): 1584 - 1595. [Full Text] [PDF]
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 Y. Suarez, B. R. Shepherd, D. A. Rao, and J. S. Pober Alloimmunity to Human Endothelial Cells Derived from Cord Blood Progenitors J. Immunol., December 1, 2007; 179(11): 7488 - 7496. [Abstract] [Full Text] [PDF]
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