Enhanced wound vascularization using a dsASCs …sites.utexas.edu › texas-bmes › files › 2015...
Transcript of Enhanced wound vascularization using a dsASCs …sites.utexas.edu › texas-bmes › files › 2015...
ORIGINAL PAPER
Enhanced wound vascularization using a dsASCs seeded FPEGscaffold
David O. Zamora • Shanmugasundaram Natesan •
Sandra Becerra • Nicole Wrice • Eunna Chung •
Laura J. Suggs • Robert J. Christy
Received: 31 January 2013 / Accepted: 29 April 2013 / Published online: 26 May 2013
� Springer Science+Business Media Dordrecht (outside the USA) 2013
Abstract The bioengineering of autologous vascular
networks is of great importance in wound healing. Adi-
pose-derived stem cells (ASCs) are of interest due to their
ability to differentiate toward various cell types, including
vascular. We hypothesized that adult human ASCs
embedded in a three-dimensional PEG-fibrin (FPEG) gel
have the ability to modulate vascularization of a healing
wound. Initial in vitro characterization of ASCs isolated
from discarded burn skin samples (dsASCs) and embedded
in FPEG gels indicated they could express such pericyte/
smooth muscle cell markers as a-smooth muscle actin,
platelet-derived growth factor receptor-b, NG2 proteogly-
can, and angiopoietin-1, suggesting that these cells could
potentially be involved in a supportive cell role (i.e., per-
icyte/mural cell) for blood vessels. Using a rat skin exci-
sion model, wounds treated with dsASCs-FPEG gels
showed earlier collagen deposition and wound remodeling
compared to vehicle FPEG treated wounds. Furthermore,
the dsASCs-seeded gels increased the number of vessels in
the wound per square millimeter by day 16 (*66.7 vs.
*36.9/mm2) in these same studies. dsASCs may support
this increase in vascularization through their trophic con-
tribution of vascular endothelial growth factor, as deter-
mined by in vitro analysis of mRNA and the protein levels.
Immunohistochemistry showed that dsASCs were localized
to the surrounding regions of large blood-perfused vessels.
Human dsASCs may play a supportive role in the forma-
tion of vascular structures in the healing wound through
direct mechanisms as well as indirect trophic effects. The
merging of autologous grafts or bioengineered composites
with the host’s vasculature is critical, and the use of
autologous dsASCs in these procedures may prove to be
therapeutic.
Keywords Angiogenesis � ASCs � Wound healing �Fibrin � Collagen � PEG
Introduction
Extremity soft-tissue loss, resulting from direct impact or
thermal injury, is common to most military conflicts. His-
torically these wounds constitute the majority of injuries,
cost burden, and tend to increase the overall length of
hospitalization [1]. During Operation Iraqi Freedom and
Operation Enduring Freedom the frequent use of impro-
vised explosive devices resulted in an increase in complex
burn injuries involving a high percentage of total body
surface area (TBSA) [2, 3]. Current treatment strategies for
such burn injuries consists of immediately covering the
debrided wound areas with autografts or acellular allografts
[4]. Autologous skin grafts have better prognosis for
Disclaimer: The opinions and assertions contained herein are the
private views of the authors and are not to be construed as official or
reflecting the views of the Department of Defense or Department of
Army. The authors are employees of the U.S. Government, and this
work was prepared as part of their official duties. This research was
funded by the U.S. Army Medical Research and Materiel Command.
This study has been conducted in compliance with the Animal
Welfare Act, the implementing Animal Welfare Regulations, and the
principles of the Guide for the Care and Use of Laboratory Animals.
D. O. Zamora � S. Natesan � S. Becerra � N. Wrice �R. J. Christy (&)
Regenerative Medicine Research Program, United States Army
Institute of Surgical Research, 3698 Chambers Pass, BHT 1:
Bldg 3611, Fort Sam Houston, TX 78234-6315, USA
e-mail: [email protected]
E. Chung � L. J. Suggs
Department of Biomedical Engineering, The University of Texas
at Austin, 1 University Station, Austin, TX 78712-0238, USA
123
Angiogenesis (2013) 16:745–757
DOI 10.1007/s10456-013-9352-y
incorporation and wound healing, but autografts create a
new wound on the patient. Furthermore, undamaged skin
tissue that can be used for grafting is limited in patients
with large TBSA burns. Therefore, treatment of large
TBSA burn patients necessitates the use of allografts skin
substitutes. Currently there are a number of commercially
available products on the market (e.g., Integra, Alloderm)
however, the function of these grafts may be compromised
due to lack of cell infiltration and tissue revascularization.
Oxygen and nutrient perfusion typically occurs within a
100- to 200-lm area of a blood vessel; any cellular com-
ponents with a distance greater than this will be exposed to
ischemic conditions and eventually undergo apoptosis.
Unfortunately, acellular grafts are limited by the length of
time it takes for cell infiltration of the tissue, inosculation
and revascularization [4–6]. Therefore, developing alter-
native strategies to deliver cells and quickly revascularize
these wounds is essential.
New bioengineered skin substitutes that deliver cells
within the graft, promote revascularization and therefore
increase long-term survival and remodeling of an allograft
skin substitute are being developed. Initial skin substitutes
used such strategies as co-culturing of vascular endothelial
cells and fibroblast cells in a collagen based matrix [7], self
assembled co-cultured human umbilical vein endothelial
cells (HUVECs) and dermal microvascular cells [8],
sequentially seeding the apical and basal surfaces of acel-
lular dermis with cultured human keratinocytes and HU-
VECs [9], and pre-seeding endothelial progenitor cells
(EC) [10]. These methodologies have had some success.
Unfortunately, obtaining vascular and/or keratinocyte cells
from a patient with large TBSA wounds, where there is a
limited source of cells to develop these substitutes is
problematic. Current strategies to overcome this problem
focus on optimizing the two main components of tissue
engineered grafts: biomaterial scaffolds and cells. The use
of stem cells, such as epidermal stem cells or mesenchymal
stem cells from bone marrow or adipose tissue, which have
the potential to differentiate into various phenotypes, rather
than using specific cell types (e.g. keratinocytes) may
provide a strategy for functional wound repair and com-
plete regeneration of skin [11]. Many different biomaterials
have been developed and shown to have beneficial effects
on wound healing and provide a microenvironment that
allows cells to proliferate and differentiate. Natural poly-
mers including the extracellular matrix proteins, collagen
and hyaluronic acid have been used extensively in wound
repair and for acellular skin grafts because of their in vitro
and in vivo cell biocompatibility [4, 12]. Fibrin, another
natural biopolymer, has been used clinically as a hemo-
static agent and as a sealant for soft tissue wounds. Still the
major concern of using fibrin-based hydrogels as wound
repair scaffolds is their relatively quick contraction, low
mechanical stiffness (which limits durability), and their
rapid degradation once placed at the wound site. These
aspects alone can hinder proper tissue formation and
reconstruction. To overcome these problems, fibrin has
been modified to increase its durability, ease of use, and
longevity in a wound. One such modification involves the
copolymerization of fibrin with polyethylene glycol (PEG)
[13, 14]. The addition of extra cross-linking between fibrin
and polyethylene glycol (FPEG) during thrombin mediated
fibrin polymerization produces a highly hydrated gel
microenvironment allowing cell seeding within the matrix
for direct delivery of cells to a wound. This combining
stem cells with the natural biological activity of fibrin has
been shown to encourage tissue and blood vessel in-growth
in the healing wound [15, 16]. Based on these criteria,
FPEG is an excellent candidate for combinatorial therapies
involving both biomaterials and stem cells.
When considering stem cells as a potential cell source
to be combined with FPEG; adipose-derived stem cells
(ASCs) are an attractive cell source due to their ease in
isolation, relative abundance, ability to differentiate
toward various cell types in vitro, and their beneficial
function in wound healing in vivo [17, 18]. For example,
ASCs have been shown to successfully enhance revascu-
larization of ischemic hind limbs in mice over such other
cells types as human mesenchymal stem cells (MSCs)
from bone marrow [19]. ASCs derived from humans
appear to assist in reestablishing blood flow in ischemic
tissue salvage experiments [20]. Intramyocardial injections
of human ASCs have been found to assist in local angio-
genesis following myocardial infarction in animals [21]. In
addition, recently it has been shown that ASCs embedded
in a dermal substitute help to improve the regeneration of
skin by increasing blood vessel formation and collagen
synthesis [22]. ASCs also have great implications in their
ability to enhance superficial wound healing. Local
implantation of ASCs has proven effective in supporting
epidermal healing in full-thickness skin wounds of pigs
and rats [23, 24].
Recent studies have shown that rat and human ASCs,
isolated from epididymal fat or from discarded human burn
tissue (dsASCs), also have plasticity to differentiate
towards vascular cell phenotypes when cultured in three-
dimensional matrices in vitro [25, 26]. We hypothesize that
the dsASCs embedded in three-dimensional gels have the
ability to differentiate into vascular cell types in vivo and
therapeutically modulate wound healing and vasculariza-
tion. In this study, we have systematically combined
human dsASCs, human fibrinogen, and polyethylene glycol
to create a unique matrix to be investigated using an
established wound healing model.
746 Angiogenesis (2013) 16:745–757
123
Materials and methods
dsASCs isolation
Discarded human burn tissues were isolated according to
our previously published protocol [25]. In brief, ASCs were
isolated from discarded skin samples from patients under-
going burn wound debridement at the U.S. Army Institute
of Surgical Research (USAISR) Burn Center, Fort Sam
Houston, Texas. The skin samples were brought to the
laboratory immediately after debridement and processed.
This study was conducted under the protocol reviewed and
approved by the U.S. Army Medical Research and Materiel
Command Institutional Review Board. Discarded burn
tissue samples were collected in accordance with the
approved protocol, HSC20080290 N. The authors were
blinded from some patient information including burn
depth, percentage of TBSA, and anatomical location of the
burn. The skin samples were washed 3–4 times with
Hank’s buffered salt solution (HBSS) to remove adherent
blood clots. The hypodermal layer was dissected away
from the dermal region, transferred to a Petri dish, and
finely minced with scissors. The minced tissue was sus-
pended in HBSS and centrifuged for 10 min at 5009g at
16 �C. The floating tissue was carefully collected; and to
every 1 ml of the floating fraction tissue, 3,500 units of
collagenase type II (Sigma-Aldrich, St. Louis, USA) was
added and incubated for 45–60 min at 37 �C in an orbital
shaker incubator at 125 rpm. The undigested tissue was
removed by sequential passage through 100- and 70-lm
nylon mesh filters. The filtrate was then centrifuged at
5009g for 10 min at 16 �C, treated with BD Pharm
LyseTM lysing buffer (BD Bioscience, San Jose, CA, USA)
to remove any remaining red blood cells, and washed twice
with HBSS. The final cell pellets were resuspended in
growth media (MesenPRO RSTM basal medium), supple-
mented with MesenPRO RSTM growth supplement, anti-
biotic–antimycotic (100 U/ml of penicillin G, 100 lg/ml of
streptomycin sulfate, and 0.25 lg/ml of amphotericin B),
and 2 mM of L-glutamine (Life Technologies, Carlsbad,
CA, USA). This medium preparation is represented as
‘‘complete medium.’’ The resulting cell number, typically
*1.7 9 106, were cultured in T75 flasks (BD Falcon, NJ,
USA) and maintained in a 5 % carbon dioxide (CO2)
humidified incubator at 37 �C. After 4 h in culture, the
growth medium was replaced to remove any floating deb-
ris. The remaining attached cells are regarded as dsASCs.
Bioscaffold preparation
Fibrin and polyethylene glycol hydrogels were prepared as
previously described by Zhang et al. [13]. Succinimidyl
glutarate modified polyethylene glycol (SG-PEG-SG;
3,400 Da; NOF America Corporation, White Plains, NY,
USA) was dissolved using 4 ml of tris-buffered saline
(TBS, pH 7.8, Sigma-Aldrich), and then sterilized using a
0.22-lm filter just prior to starting the experiment to obtain
a stock solution of 8 mg/ml. Next, 500 ll of fibrinogen
stock (40 mg/ml in TBS) and 250 ll of PEG stock (8 mg/
ml) were mixed in a 12-well cell culture plate and incu-
bated for 20 min in a 5 % CO2 humidified incubator at
37 �C. This mixture constitutes a molar concentration ratio
of 10:1, SG-PEG-SG: fibrinogen. After incubation, 250 ll
of dsASCs (100,000 cells in MesenPRO complete medium)
were mixed with the PEGylated fibrinogen solution, and
immediately 1 ml of thrombin stock (25 U/ml; Sigma-
Aldrich) in 40 mM of calcium chloride (CaCl2) at a final
concentration of 10 U/ml was added. The 2-ml solution
was then quickly triturated once with the pipettor, and
immediately 1-ml aliquots were placed individually in a
12-well format cell culture insert of 8-lm pore size. Once
gel-cell mixtures were aliquoted into their respective wells,
the mixtures were incubated in a 5 % CO2 humidified
incubator at 37 �C for 10 min to allow for complete gela-
tion. The resulting FPEG gels were then washed twice with
HBSS and incubated with alpha minimal essential media
(a-MEM) supplemented with 10 % FBS in a 5 % CO2
humidified incubator at 37 �C. The formation of tube-like
networks by dsASCs migration was then observed over an
11-day period using standard light microscopy techniques.
RT-PCR analysis
Total ribonucleic acid (RNA) from dsASCs in FPEG gels
(3, 5, 7, 9, and 11 days) were isolated using TRIzol� LS
Reagent (Invitrogen), with slight modifications. Before
processing, the gels were rinsed with HBSS once and
carefully removed from the culture well. Four gels from
each time point were pooled together and minced, 16 ml of
TRIzol� LS Reagent was added, and then samples were
incubated for 15 min on ice. After incubation, 8 ml of
chloroform was added, samples were mixed, and the
aqueous phase was separated by centrifugation
(13,0009g). Total RNA was then purified using Qiagen’s
mini spin columns (Qiagen, Valencia, CA, USA). The
concentration and quality of the purified RNA was deter-
mined at 260/280 optical density ratio using a NanoDrop
spectrometer (NanoDrop Technologies, Inc., Wilmington,
DE, USA). Complementary deoxyribonucleic acid (cDNA)
was synthesized from 150 ng of total RNA, using Super-
Script III first-strand synthesis supermix with oligo-dT
primers (Invitrogen). A negative control sample (H2O) was
used to assess random production of cDNA through con-
taminants. Oligonucleotide primer sequences (Acta2,
CD140b, NG2, Angpt-2, Angpt-1, and VEGF165) were
purchased from SA Biosciences (A Qiagen Company;
Angiogenesis (2013) 16:745–757 747
123
Frederick, MD). Master mixes were made containing 200
nM of forward and reverse primers containing SYBR
Green-ER, and qPCR supermix (Invitrogen); and the syn-
thesized cDNA was added to the appropriate wells. Real-
time polymerase chain reaction (RT-PCR) was carried out
using a Bio-Rad CFX96 thermal cycler system (Bio-Rad,
Hercules, CA, USA). Message RNA expression levels were
normalized to glyceraldehye-3-phosphate dehydrogenase
(GAPDH). Fold increase in expression levels for each
specific gene was normalized to the expression levels of
control passage 2 dsASCs. Fold increase in expression
levels for each gene was determined by the 2-DDCt method.
Immunochemistry
To analyze the differentiation of dsASCs, cells were seeded
within the FPEG gels and allowed to grow for 11 days
before being harvested. On the day of harvest, the medium
was aspirated from the wells; the FPEG-dsASCs gels were
gently removed from their cell culture inserts using a
spatula and immediately processed for immunocytochem-
istry. Briefly, the gels were washed with HBSS (twice,
5 min), fixed with 4 % paraformaldehyde (EMS, Hatfield,
PA) for 20 min, treated serially with increasing concen-
trations of sucrose (from 5 to 20 %, 30 min each incuba-
tion), and then incubated overnight with 20 % sucrose at
4 �C. The sucrose-treated gels were then embedded in a
20 % sucrose-HistoPrep (Fisher Scientific, Pittsburgh, PA,
USA) mixture (2:1) and flash-frozen using liquid nitrogen.
Sections, 10–12 lm thick, were then cut using a cryostat
(Leica Microsystems, Nussloch, Germany), washed with
sterile HBSS, and fixed with 4 % paraformaldehyde for
20 min. Nonspecific Fc receptor-mediated sites were
blocked by incubating the sections for 1 h with 1 % bovine
serum albumin in HBSS containing 0.01 % Triton X-100
and then washed twice (5 min) with HBSS. To assess the
endothelial immunophenotype, sections were stained with
anti-human CD31 (PECAM-1, 8 lg/ml; R&D Systems)
and von Willebrand factor (vWF, 10 lg/ml; Millipore,
Billerica, MA, USA) specific monoclonal primary anti-
bodies. For identifying pericyte immunophenotype,
human-specific monoclonal antibodies specific to chron-
droitin sulfate proteoglycan (NG2, 20 lg/ml; Millipore),
and platelet-derived growth factor receptor beta (PDGFRb/
CD140b, 10 lg/ml; R&D Systems) antibodies were used.
After incubation of unconjugated primary labeled anti-
bodies, sections were washed twice (5 min) with HBSS
and incubated with 5 lg/ml host species-specific Alexa
fluor 488 and/or Alexa fluor 594 secondary antibodies
(Invitrogen) for 45 min at 4 �C. Finally, the sections were
washed twice with HBSS (5 min) and nuclei stained with
Hoechst 33342 (Invitrogen). Nonspecific fluorescence was
evaluated using sections incubated with respective
fluorophore-labeled secondary antibodies. Epifluorescence
of cells and gel sections were observed using Olympus
IX71 inverted microscope equipped with reflected fluo-
rescence system (Olympus America Inc.). Photomicro-
graphs were taken using a DP71 digital camera, and image
overlay was carried out using DP controller application
software.
Animal model
Male Rowett nude (RNU) rats (athymic rats), deficient in T
cell function, weighing 175–250 g, were obtained from
Harlan Laboratories (Indianapolis, IN) and housed in the
animal care facility at the USAISR with access to water
and rat chow ad libitum. This study was conducted in
compliance with the Animal Welfare Act, the implement-
ing Animal Welfare Regulations, and the principles of the
Guide for the Care and Use of Laboratory Animals. The
rats underwent general anesthesia for the surgery after
receiving a preemptive dose of analgesic (buprenorphine
0.1 mg/kg, subcutaneous) 30 min prior to induction.
Anesthesia was induced by placing rats into a plexiglass
chamber filled with 1–3 % isoflurane and oxygen. Anes-
thetic was maintained using a vaporizer setting of 1–3 %
isoflurane delivered with a nose cone on a Bain circuit
hooked to the rodent gas anesthesia machine (VetEquip,
Inc., Pleasanton, CA, USA). A full-thickness skin excision
wound 1.5 cm in diameter was created on the dorsum of
the rat down to the panniculus. The rats were randomly
divided into three groups with a total of two rats per group
and treated as follows: saline control group (250 ll of
saline), FPEG gel, and FPEG-dsASCs gel treatments. Upon
placement of treatments, the wounds were covered with
DuoDERM� dressing (ConvaTec, Skillman, NJ, USA) and
evaluated at 4, 8, 12, and 16 days. When the study was
completed, the rats were euthanized; and the wound beds,
including the healthy skin margin of the healed area sur-
rounding the wound, were harvested and fixed with 10 %
neutral-buffered formalin for histological analysis.
Histology and vessel quantification
Histological analysis was performed on *5 lm sections of
formalin-fixed paraffin embedded granulation tissue and
normal skin tissue surrounding the wound collected from
the wound bed of the athymic nude rats. The sections were
stained with either hematoxylin and eosin or Masson’s
trichrome and examined under light microscopy to assess
the overall wound healing pattern in the rat tissue. For
blood vessel identification, paraffin embedded tissues were
immunolabeled with anti-vWF antibody (Cat#250A-1,
rabbit polyclonal) using Cell Marque (Rocklin, CA, USA)
and the Ventana automated tissue immunostaining unit
748 Angiogenesis (2013) 16:745–757
123
located at the Brooke Army Medical Center Histology
Laboratory, Fort Sam Houston, Texas. Prepared slides
were then analyzed using standard light microscopy and
blood vessels [*10 lm in luminal diameter were quan-
titated in the center of the wound. The parameters that
qualified a vessel to be counted are as follows: (1) positive
vWF staining, (2) vessels were counted only once even if
multiple sections of the same vessel appeared in the field of
view, as assessed by trajectory of vessel and diameter size
and (3) vessel had to be located within a 1.5-cm window
centrally located within the wound, as delineated by the
Olympus microscope software. Vessels quantified were
located within the newly forming tissue-gel treatments,
taking care to avoid vessels in the host tissue below and
surrounding the regenerating wound. For any given slide,
five 1-mm2 regions located within 1.5-cm wound center
were counted in a blinded fashion. The five regions counted
were averaged and are represented in Fig. 6a as raw data
per animal. To analyze the diameter of vessels that the
donated human cells associated with, dual labeled vessels
for human specific mitochondrial antigen (h-MT) and
CD31 were digitally measured using the Olympus micro-
scope software in vessels of these same regions.
Enzyme-linked immunosorbent assay (ELISA)
To assess VEGF protein production by dsASCs, the cells
(P2) were seeded within the FPEG gels similarly to the gels
prepared for the animal studies (50,000 cells/ml of gel).
However, in these experiments, the gels were cast within a
6-well plate filter insert (8.0 lm pore size; BD Falcon,
Franklin Lakes, NJ, USA). FPEG gels without dsASCs
were processed simultaneously as controls. Three millili-
ters of complete medium was added to the bottom chamber
and 2 ml of complete medium to the top chamber of the
filter insert, and then plates were placed in a 5 % CO2
humidified incubator at 37 �C. A complete medium change
was performed daily on both the inner and the outer wells.
Prior to removing the medium, 1 ml of conditioned med-
ium was collected from each sample and stored at -80 �C
until ready for analysis. Conditioned medium from the
corresponding time points (1, 3, 5, 7, 9, 11, and 15 days) were
quantified for the levels of human VEGF165 using com-
mercial ELISA kits (Quantikine�, R&D Systems) per
provided protocols and standard curves.
In vivo dsASCs localization
Cryosections (*10 lm) were cut from frozen tissue sam-
ples using a Leica CM1850 (Leica Microsystems, Buffalo
Grove, IL, USA) at a set temperature of -20 �C. Sections
were applied to poly-prep slides (Sigma-Aldrich, St. Louis,
MO, USA) and stored at -80 �C until further processed.
Tissue sections were acclimated to room temperature for
20 min and then rehydrated with 19 wash buffer (Dako,
Carpinteria, CA, USA) for 10 min. The endogenous
enzymes were blocked according to EnVision G/2 Dou-
blestain System (Dako), rabbit/mouse (diaminobenzi-
dine ? permanent red) kit instructions. To localize human-
specific cells within the tissue, sections were incubated with
10 lg/ml of human-specific anti-mitochondrial (h-MT),
mouse-purified monoclonal primary antibody (10 lg/ml,
Millipore, Billerica, MA, USA) for 1 h at room temperature
followed by an alkaline phosphatase/horseradish peroxi-
dase-labeled secondary antibody polymer system. Primary
labeled tissue was further processed for dual labeling fol-
lowing instructions from the labeling kit (Dako). Sections
were then incubated with 5 lg/ml of second primary anti-
body for 2 h using anti-platelet endothelial cell adhesion
molecule (PECAM-1; Millipore) to identify blood vessel
formation within the tissue. Concentration requirements for
the second primary antibody, anti-PECAM, was incubated
for 2 h at room temperature. Close attention was paid when
developing the tissue with diaminobenzidine and permanent
red chromagen because the rate varies with tissue type.
Lastly, some tissues were counterstained with 0.2 % methyl
green (Vector Laboratories, Burlingame, CA, USA) for
10 min to stain nuclei, then air-dried and mounted using an
aqueous mounting medium.
Results
Differentiation of dsASCs
Previous studies by our lab fully characterized the pheno-
type of the isolated dsASCs used in this study and con-
firmed the stem cell nature of these cells [25]. To investigate
the potential of the dsASCs (Fig. 1a) to differentiate toward
vascular phenotype lineages, the cells were cultured in a
three-dimensional FPEG gel (Fig. 1b) and maintained over
an 11-day period and analyzed at different time points (3, 5,
7, 9, and 11). Within 3 days of culture (Fig. 1c), the cells
had began sprouting tube-like structures; and by days 7 and
11 they became denser (Fig. 1d, e, respectively). Previous
reports by our lab [26] and others [27, 28] have demon-
strated that rat ASCs have an ability to express an endo-
thelial cell phenotype. Since the tube-like structures that the
human dsASCs formed within the FPEG were reminiscent
of endothelial cell tube structures, we analyzed these cul-
tures for their expression of CD31 and vWF. Interestingly,
CD31 and vWF were undetectable in these cultures by our
immunohistochemistry and PCR assays at the time points
indicated (data not shown). Current dogma indicates that
ASCs may exist in vivo as a pericytic niche [29]; therefore,
we investigated their potential for pericyte marker
Angiogenesis (2013) 16:745–757 749
123
expression (Fig. 1f). In these three-dimensional culture
conditions, dsASCs were induced to express pericyte-spe-
cific markers when cultured in two-dimensional tissue
culture conditions, as determined by 2-DDCt. A slight
increase in expression (*1–5-fold) of smooth muscle a-
actin (Acta2) and CD140b was observed, with little mod-
ulation of their expression over an 11 day period. A modest
increase (*15 fold by day 11) in NG2 expression was
observed over this same timeframe. Angpt-1 transcript,
which is expressed mainly by perivascular cells and mural
cells, increased in these cultures between days 3 and 7 and
returned to baseline levels by day 11, perhaps indicating the
termination of network formation. Angpt-2, a negative
regulator of Angpt-1 signaling during angiogenesis is gen-
erally known to be expressed by endothelial cells and
mesenchymal stem cells. In these cultures, the cells
exhibited a substantial fold increase (*5- to 15-fold
increase) of Angpt-2 between days 3 and 9 but drastically
Fig. 1 Molecular and
morphological characterization
of human dsASCs. a Phase-
contrast photomicrograph
depicting the morphology of
dsASCs (P1), b image depicts
the *1.5-cm-diameter FPEG
gel after removal from the filter
insert. The orange rectangle
depicts the gel at *0.3- to 0.5-
cm thickness. Once seeded in
the FPEG gel, dsASCs exhibited
a tubular network formation
over time c day 3, d day 7, and
e day 11. f Total RNA was
isolated from dsASCs and
FPEG-dsASCs gels (days 3–11)
and analyzed by real-time PCR
for transcript expression of
endothelial (CD31, vWF) and
pericytic (ACTA2, CD140b,
NG2, Angpt-1 and Angpt-2)
markers. While dsASCs
demonstrated mRNA expression
of pericytic markers over the
time frames examined (shown),
endothelial markers remained
undetectable in our assays (data
not shown). Gene expression
levels are represented as mean
fold changes (±standard
deviation). (g-l)
Immunostaining of dsASCs
cultured in FPEG gels for
11 days confirmed RT-PCR
results demonstrating the
expression of such pericytic
markers by dsASCs. Original
magnifications: a, c, d, e 9200;
g–l: 9400. (Color figure online)
750 Angiogenesis (2013) 16:745–757
123
down-regulated by day 11. Initial expression of Angpt-2
may be indicative of the inherent ‘‘stemness’’ of these cells.
To corroborate PCR data, protein expression of CD31,
vWF, and CD140b and NG2 markers by the cells was
examined by immunofluorescence staining. Similarly, cell
cultures did not stain positive for vascular endothelial
markers CD31 and vWF (data not shown) but did stain
strongly for NG2 and CD140b (Fig. 1h, k, respectively).
Wound healing and vascularization
To investigate the influence of FPEG-dsASCs gels on
wound healing, we implemented our established rat full-
thickness skin excision wound model and treated the
wounds with either dsASCs seeded FPEG gels, FPEG gels
alone, or saline, Fig. 2a (i–iv). The treatments were left on
the wounds over 4, 8, 12, and 16 days. At these time points,
the rats were euthanized; and wound beds, along with
normal skin adjacent to it, were examined by histological
technique. Overall, wounds appeared to close at similar
rates between all treatment groups. However, wound
remodeling and collagen deposition (blue stain in Figs. 2b,
4b) appeared to occur sooner in FPEG-dsASCs treated
wounds than in control treatments. Closer examination
showed that vessel-like structures were also present and
that these structures appeared sooner in the FPEG-dsASCs
treated wounds (day 8; Fig. 3) than in wounds lacking cell
treatment. The vessel-like structures in Figs. 3c, 4a, b did
contain red blood cells and were lined with a monolayer of
cells. To confirm that these vessel-like structures are per-
fused blood vessels, sections were immunostained for the
vascular endothelial cell specific marker vWF. The sections
showed positive vWF staining confining to the vessel-like
structures, thus confirming the vascularization of the
wound bed (Fig. 5). Upon quantitating blood vessel den-
sity, it became apparent that FPEG-dsASCs treatment
increased the amount of blood vessels in the healing
wound, compared to FPEG alone and saline treatments
(Fig. 6a). Furthermore, the blood vessels within the wound
bed of FPEG-dsASCs treatments appeared to be larger and
stained darker for vWF (Fig. 6b, c) than FPEG treatments
alone (Fig. 6d, e) and suggested that the presence of
dsASCs may enhance angiogenesis in these tissue gels.
Role of the dsASCs
To determine whether dsASCs play a direct role in blood
vessel formation in vivo, we needed to localize the dsASCs
within the wound bed with a h-MT specific antibody. With
this reagent, the human cells consistently localized to
medium sized vessels of *15.06 lm diameter, but not
smaller capillary vessels *5.27 lm, (Fig. 7a, d). It is
Fig. 2 Gross histology of treated excision wound. a (i) Circular
excision wound created on the dorsum of athymic nude rats. a (ii)
Clear FPEG gel, ±dsASCs, is immediately placed into the excised
wounds and securely bandaged into place. a (iii) Photomicrograph of
a day 4 excised wound bed that was cross-sectioned mid-sagitally.
Higher magnification of white rectangle is depicted in a (iv). The
horizontal dashed line in a (iii, iv) demarcates the host wound tissue
from the FPEG gel. Above the dashed line is the FPEG gel; below the
dashed line is the host. Gross examination of the wound bed indicated
that the groups treated with FPEG gels ± dsASCs had completely
integrated with the host tissue by day 12 (b). Masson’s trichrome-
stained tissue sections from all treatment groups on days 4, 8, 12, and
16. Original magnifications: a Gross photograph; b 940
Angiogenesis (2013) 16:745–757 751
123
important to note that FPEG gel treatments alone contained
little to no medium sized vessels, but did have smaller
capillary vessels that ranged in diameter from 4 to 9 lm.
This is consistent other analysis depicted in Figs. 3b, 5
(Days 8 & 16), and 6d, e. To analyze this observation,
higher magnification of the vessels indicated that the
human cells were incorporated into the vessel structure by
physically surrounding the blood vessel in a similar fashion
that pericytes do (Fig. 7b; brown stain) but did not appear
to localize to the inner CD31? endothelial lining (red
stain). Anti-CD31 antibodies used were cross-reactive to
both rat and human species alike.
To investigate a potential mechanism for dsASCs to
increase blood vessel density in a healing wound, we
examined their potential for VEGF expression in vitro
within a 3D FPEG gel culture system. FPEG-dsASCs gels
were harvested at different time points (3, 5, 7, 9, 11, and
15 days), and total RNA was isolated and analyzed for
VEGF expression. As early as 3 days after culture, an
increase in their VEGF expression was observed, as com-
pared to dsASCs cultured in a standard two-dimensional
culture system (Fig. 8a). A constant increase in the
amounts of VEGF mRNA expression by dsASCs was
observed over the period of culture. This observation also
translated into soluble VEGF protein by these cells, as was
determined by ELISAs of conditioned media during the
same time points (Fig. 8b).
Discussion
Extremity trauma from blunt force or penetrating objects
and chemical or thermal burns remains a leading cause of
morbidity for civilian and military personnel alike. More-
over, the healing and tissue regeneration of such large-
volume wounds remains a significant hurdle in treating
these patients and restoring functionality to their limbs. As
a part of the treatment regimen, necrotic or burned tissue is
surgically debrided from the wound site and discarded.
Often during this process, viable tissue is inevitably
removed as well. We previously reported on the ability to
isolate ASCs from the viable hypodermis region of deb-
rided skin (dsASCs), which is typically removed from burn
injured patients [25]. These cells are able to be isolated in
significant quantities for clinical use and maintain their
stem cell marker expression in vitro. In the present study,
we extend these findings by investigating their therapeutic
use in the angiogenic process of wound healing and tissue
regeneration when used in combination with a novel FPEG
hydrogel.
ASCs and vascularization
The notion that ASCs could be beneficial for the regener-
ation of vascular systems, either through endothelial or
mural cell differentiation or the secretion of paracrine fac-
tors, has been well documented in the literature over the
past decade [18, 23, 30–33]. In particular, attention has been
focused on the ability of ASCs to differentiate toward
vascular endothelium [26–28]. A number of reports indicate
that both human and animal ASCs can molecularly and
phenotypically resemble endothelium, but only up to a
certain extent. Generally, exogenous growth factors (i.e.,
VEGF, EGF) are supplemented into culture conditions to
induce ASCs toward EC differentiation. However, these
cells do not appear to fully express, simultaneously, all
characteristics of an end-differentiated endothelial cell (i.e.,
CD31, vWF, or eNOS expression, LDL uptake, single-layer
cobble-stone morphology, or tube formation on Matrigel).
Early studies by our lab and colleagues using rat ASCs
indicated that these cells were capable of expressing such
endothelial markers as CD31 and vWF, as well as form
lumen containing capillary-like tubes in vitro. Moreover,
this was accomplished without the use of growth factors and
was attributed to the unique composition of the FPEG 3D
hydrogel. Using similar methodologies, human dsASCs
Fig. 3 Histology of treated wounds. a, b, c Day 8 wounds were more
closely examined as both b FPEG and c FPEG-dsASCs wounds
appeared to possess functional blood vessels. Closer examination
indicated that wounds treated with FPEG-dsASCs had numerous
organized vessel structures (hashed oval region), whereas FPEG gel
(b) or saline (a) treatments alone exhibited fewer blood vessels.
Original magnifications: 9100
752 Angiogenesis (2013) 16:745–757
123
were isolated and their vasculogenic potential investigated
here. Initial studies indicated that these human dsASCs
were capable of forming networks in the three-dimensional
FPEG gel (Fig. 1). However, under these conditions,
dsASCs did not appear to express CD31 or vWF (data not
shown) but did express such pericytic markers as smooth
muscle actin (Acta2), platelet-derived growth factor
receptor beta (CD140b), chrondroitin-sulfate proteoglycan
(NG2), and angiopoietin (Angpt-1). Furthermore, dsASCs
appeared to target and surround newly formed blood vessels
of approximately 15.06 lm diameter, in a rat skin excision
wound model, but did not appear to localize at the inner
endothelial lining of the vessels. At no time were human
dsASCs detected as being associated with vessels of smaller
diameter (*8 lm or less). Whether this is reflective of
actual biology or beyond the limit of our detection still
remains to be determined. The immunolabeling method
used here is a unique technique that is specific for the
localization of human-specific mitochondrial antigen in
human, not rat, cells. Taken together, this data suggests that
the implanted human dsASCs appear to differentiate and
function in a supportive pericytic role and not as a lumenal
endothelial cell, as observed in our animal model. This
observation is in line with the emerging hypothesis that
ASCs, a subtype of mesenchymal stem cells, have a peri-
cytic niche and localize to perivascular compartments to
support and stabilize blood vessels. A current model pro-
posed by da Silva et al. [29] is that the ASCs-pericytic niche
stabilizes blood vessels and contributes to tissue and
immune system homeostasis under physiological conditions
but is able to assume more of an active tissue regeneration
role upon injury. In our animal model, it appears that the
dsASCs survive the transplant and home to the perivascular
region of the newly formed blood vessels.
ASCs and their paracrine effects
ASCs secrete a variety of growth factors that can exert their
effects on wound repair and tissue regeneration. Some of
these factors include basic fibroblast growth factor, hepa-
tocyte growth factor, insulin growth factor binding protein,
granulocyte colony-stimulating factor, and transforming
growth factor beta 1 [31, 32]. The potential to express such
a vast cytokine profile makes ASCs attractive not only for
their ability to differentiate toward other cells types but
also for their ability to therapeutically affect wound healing
by the secretion of such paracrine factors. It is noteworthy
to mention that such factors also have the ability to mod-
ulate immune and inflammatory responses at the wound
site, recruit local stem cells into the wound microenvi-
ronment, and reduce the amount of apoptotic cells, all
while promoting an angiogenic response [18]. All of these
effects are desirable for creating a microenvironment that is
conducive for tissue regeneration. VEGF is also secreted
by ASCs and is mainly recognized for its ability to promote
vascular endothelial cell proliferation and migration [33].
However, VEGF adds a multitude of benefits to a healing
wound, aside from endothelial cell recruitment, that should
be further considered in a wound healing scenario. Studies
have shown that VEGF is sufficient to induce fibroblast
migration, which is a critical component of the wound-
healing process. VEGF is also known to increase the epi-
thelialization of healing dermal wounds [34]. It is worth
noting that other isoforms of VEGF exist and, in particular,
that VEGF-E also induces keratinocyte migration and
proliferation [35]. It remains to be determined whether
ASCs express all forms of VEGF or just VEGF-A. In our
studies, ASCs expressed VEGF-A transcript when cultured
Fig. 4 Functional blood vessels in the regenerated tissue. a Closer
examination of FPEG-dsASCs treated tissues (day 8) indicated well
formed blood vessels within the regenerating tissue, complete with
endothelium (arrows) and red blood cells within the lumens.
Fibroblast-like cells (dark pink), oriented in perpendicular fashion
to the vessel, appeared throughout the regenerating tissue. b By day
12, collagen layers had been remodeled (blue) into mature bundles,
and function blood vessels were still present (arrow). Original
magnifications: 9400. (Color figure online)
Angiogenesis (2013) 16:745–757 753
123
Fig. 5 von Willebrand Factor (vWF) labeled vessels. To definitively
discern blood vessels in each of the treatment conditions, tissues were
immunolabeled for the vascular endothelial-specific marker vWF
(brown stain) and counterstained for contrast with hematoxylin (blue).
By day 4, both FPEG-dsASCs and FPEG gels alone appeared to have
an initial infiltration of polymorphonuclear cells (dark blue) located
within the upper portion of the gel (hashed oval). By day 8, vWF
expressing endothelium (brown stain) lined the newly formed blood
vessels in the remodeled FPEG-ASCs treated wound and, to a lesser
extent, in the FPEG-alone treated wound. By day 16, a clear difference
was observable in the amount of vessels staining positive for vWF in
the FPEG-ASCs treated group, compared to the FPEG gel alone. The
black hashed line indicates host-gel interface region, with the host
tissue below the line and remodeled gel above. The white hashed line
indicates the newly re-epithelialized region (above) with the treated
regions below. Original magnifications: 9100. (Color figure online)
Fig. 6 Vascular quantification. a Blood vessels were quantitated in
each of the treatment groups by counting the vWF? blood vessels in
sections of the healing tissues. Raw data is presented as the average
number of blood vessels counted per square millimeter in each subject
per treatment (±standard deviation). The number of vWF? vascular
structures increased over a 16-day treatment period. However, FPEG-
dsASCs treatment groups demonstrated a higher density of blood
vessels per square millimeter than FPEG treatments alone. Upon
histological examination, vessels in the FPEG-dsASCs treated groups
(b) also appeared to be larger in size than those in FPEG alone
treatment groups (d) at day 8; this trend continued through day 16 (c,
e, respectively). Special considerations were made to ensure the
accuracy of our blood vessel counts. For example, the center of the
healing wound was chosen for histological analysis to quantify blood
vessels in our model. Peripheral wound blood vessels infiltrate too
quickly to discern a difference in vessel formation, and the center of
the wound is typically the most difficult and final region of a wound to
heal. Therefore, vessel formation in this central area is more relevant
for tissue regeneration purposes than the peripheral vascular-rich
region. Original magnifications: 9400
754 Angiogenesis (2013) 16:745–757
123
in our three-dimensional FPEG gel system, and this
expression increased over time. This observation was also
true for VEGF-A protein levels being secreted into the
culture medium as well, demonstrating that the dsASCs
embedded within the gel have the ability to express VEGF-
A and that this growth factor is able to diffuse from the gel
to increase its concentration in the microenvironment of the
gel system. It remains to be determined whether this
Fig. 7 Localization of human dsASCs in wound model. a To
determine the fate of the transplanted dsASCs as the wound tissue
is remodeled, an antibody specific for human mitochondrial protein
was used to immunolabel tissue sections of FPEG-dsASCs treated
wound beds (Day 16). Human dsASCs were localized to the
surrounding regions of larger blood vessels (brown stain, solid
arrow). Interestingly, human dsASCs did not colocalize to the smaller
capillary-like vessels (hashed oval). To confirm the structures that
dsASCs associated with are indeed blood vessels, tissues were
double-labeled for PECAM-1/CD31 (red stain, hashed arrow).
b Higher magnification clearly demonstrates the red staining of
PECAM-1 along the inner luminal endothelial layer (hashed arrow),
whereas the outer brown (solid arrow) demonstrates the location of
the human dsASCs. c Photomicrograph depicting antibody isotype
controls for the immunostaining procedure. d Bar graph depicting
representative diameter sizes of dsASCs-FPEG gel treatments.
Original magnifications: a, c 9100; b 9400. (Color figure online)
Fig. 8 In vitro VEGF Expression by dsASCs. a Using real-time PCR,
VEGF transcript expression was assessed in dsASCs (P1–P4) seeded
within FPEG gels. Fold increases in transcript expression levels for
VEGF were determined by 2-DDCt method (±standard deviation), and
comparison RNA was obtained from dsASCs that were cultured side
by side under simultaneous conditions in a T25 flask. Interestingly,
when embedded within the FPEG gel, the dsASCs upregulate their
expression levels of VEGF mRNA; this observation translates into
b VEGF protein expression of VEGF as well. Each condition was
performed in triplicate, with a total three separate experiments being
performed (±standard deviation)
Angiogenesis (2013) 16:745–757 755
123
increase in VEGF protein is reflective of increasing num-
bers of ASCs due to proliferation or whether the existing
ASCs are simply expressing a higher level of the gene
encoding VEGF. Either phenomenon is favorable in this
wound-healing scenario. Equally important to VEGF
expression is the angiopoietin system. Indeed, Angpt-1 is
important in vessel stability and has been shown to con-
tribute to the formation of patent vessels of wider diameter
in vivo than control conditions [36, 37]. The synergistic
coordinated expression of VEGF and Angpt-1 by donated
dsASCs in our gel system provides a favorable pro-
angiogenic milieu of growth factors necessary for thera-
peutic angiogenesis to occur in this gel system.
FPEG and wound healing
In the clinic, fibrin has been used as an FDA-approved
hemostatic and sealant agent in a variety of applications.
Additionally, fibrin based hydrogels produced from com-
mercially available purified fibrinogen and thrombin have
been used widely in novel tissue engineering applications,
and include the engineering of such tissues as adipose,
cardiovascular, ocular, muscle, and skin tissues [38]. Such
fibrin hydrogels have also been used to promote angio-
genesis when needed [39]. Our preliminary work has
demonstrated several unique features of FPEG hydrogels
that make it attractive in wound healing over other types of
hydrogel dressings. FPEG exhibits unique features of both
synthetic hydrogels and natural materials. Studies by Ah-
mann et al. [15] showed that the degradation products of
fibrin are bioactive and can enhance vascular smooth
muscle cell proliferation and increase collagen matrix
deposition, both desirable effects in wound healing. In our
studies, the FPEG gels (±dsASCs) appeared to support a
denser and more organized collagen wound bed than saline
controls alone (Fig. 2b, day 16). Finally, fibrin possesses an
inherent biologic capability to encourage wound healing by
stimulating tissue and blood vessel in-growth [16, 40]. In
our studies described here, we observed an increase in the
presence of blood vessels in FPEG-treated wounds over
saline controls (Fig. 3), indicating that the FPEG gels alone
had beneficial effects to the healing wound. Moreover,
blood vessel formation was enhanced in FPEG-dsASCs-
seeded gels over FPEG gels alone, indicating a desirable
effect that is presumably attributed to the presence of the
dsASCs in the gel. It is possible that the enhanced blood
vessel formation in the dsASCs-FPEG treated wounds may
be a direct result of the increased VEGF production by the
dsASCs in the gel. However, since VEGF expression was
assayed in vitro, it is difficult to say with certainty that the
FPEG-dsASCs gel would perform similarly in vivo.
Nonetheless, patent red blood cell-containing vessels were
present in higher numbers than FPEG or saline-treated
wounds. It is likely that in addition to VEGF, the dsASCs
are secreting other biologic factors that influence this
enhanced vascularization. It is also likely that fibrin deg-
radation products may be playing a role in the wound-
healing process, and whether dsASCs are contributing to
the FPEG degradation remains to be determined.
Conclusion
We have successfully isolated adipose-derived stem cells
from discarded human tissue; these cells maintain their
expression profile of key stem cell markers when cultured
in vitro. Our lab has previously differentiated these cells
toward osteocytes and adipocytes, and now pericyte-like
cells. In a rat excision wound model, human dsASCs appear
to integrate with newly formed host tissue and assist to
increase the presence of functional vascular networks in this
region. Future studies will investigate the combinatorial
effect of these cells with the initial stromal vascular fraction
isolated to investigate its potential use in autologous tissue
transplants, wound repair, and skin regeneration.
Acknowledgments Dr. Natesan is supported by a Postdoctoral
Fellowship Grant from the Pittsburgh Tissue Engineering Initiative
(PTEI). Funding for this work was provided by the TATRC Foun-
dation and the Deployment Related Medical Research Program.
Conflict of interest The opinions or assertions contained herein are
the private views of RJC and are not to be construed as official or as
reflecting the views of the Department of the Army or the Department
of Defense.
References
1. Chan RK et al (2012) Development of a vascularized skin con-
struct using adipose-derived stem cells from debrided burned
skin. Stem Cells Int 2012:841203
2. Owens BD et al (2008) Combat wounds in operation Iraqi Free-
dom and operation Enduring Freedom. J Trauma 64(2):295–299
3. Wolf SE et al (2006) Comparison between civilian burns and
combat burns from Operation Iraqi Freedom and Operation
Enduring Freedom. Ann Surg 243(6):786–792; discussion 792-5
4. Brusselaers N et al (2010) Skin replacement in burn wounds.
J Trauma 68(2):490–501
5. Carmeliet P, Jain RK (2000) Angiogenesis in cancer and other
diseases. Nature 407(6801):249–257
6. Rouwkema J, Rivron NC, van Blitterswijk CA (2008) Vascu-
larization in tissue engineering. Trends Biotechnol 26(8):434–441
7. Sahota PS et al (2003) Development of a reconstructed human
skin model for angiogenesis. Wound Repair Regen 11(4):275–284
8. Sorrell JM, Baber MA, Caplan AI (2007) A self-assembled
fibroblast-endothelial cell co-culture system that supports in vitro
vasculogenesis by both human umbilical vein endothelial cells
and human dermal microvascular endothelial cells. Cells Tissues
Organs 186(3):157–168
9. Schechner JS et al (2003) Engraftment of a vascularized human
skin equivalent. FASEB J 17(15):2250–2256
756 Angiogenesis (2013) 16:745–757
123
10. Shepherd BR et al (2006) Vascularization and engraftment of a
human skin substitute using circulating progenitor cell-derived
endothelial cells. FASEB J 20(10):1739–1741
11. Zhang CP, Fu XB (2008) Therapeutic potential of stem cells in
skin repair and regeneration. Chin J Traumatol 11(4):209–221
12. Girandon L et al (2011) In vitro models for adipose tissue engi-
neering with adipose-derived stem cells using different scaffolds
of natural origin. Folia Biol (Praha) 57(2):47–56
13. Zhang G et al (2006) A PEGylated fibrin patch for mesenchymal
stem cell delivery. Tissue Eng 12(1):9–19
14. Liu H, Collins SF, Suggs LJ (2006) Three-dimensional culture for
expansion and differentiation of mouse embryonic stem cells.
Biomaterials 27(36):6004–6014
15. Ahmann KA et al (2010) Fibrin degradation enhances vascular
smooth muscle cell proliferation and matrix deposition in fibrin-
based tissue constructs fabricated in vitro. Tissue Eng Part A
16(10):3261–3270
16. Chalupowicz DG et al (1995) Fibrin II induces endothelial cell
capillary tube formation. J Cell Biol 130(1):207–215
17. Zuk PA et al (2002) Human adipose tissue is a source of multi-
potent stem cells. Mol Biol Cell 13(12):4279–4295
18. Hong SJ, Traktuev DO, March KL (2010) Therapeutic potential
of adipose-derived stem cells in vascular growth and tissue repair.
Curr Opin Organ Transplant 15(1):86–91
19. Kim Y et al (2007) Direct comparison of human mesenchymal
stem cells derived from adipose tissues and bone marrow in
mediating neovascularization in response to vascular ischemia.
Cell Physiol Biochem 20(6):867–876
20. Moon MH et al (2006) Human adipose tissue-derived mesen-
chymal stem cells improve postnatal neovascularization in a
mouse model of hindlimb ischemia. Cell Physiol Biochem
17(5–6):279–290
21. Kondo K et al (2009) Implantation of adipose-derived regener-
ative cells enhances ischemia-induced angiogenesis. Arterioscler
Thromb Vasc Biol 29(1):61–66
22. Meruane MA, Rojas M, Marcelain K (2012) The use of adipose
tissue-derived stem cells within a dermal substitute improves skin
regeneration by increasing neoangiogenesis and collagen syn-
thesis. Plast Reconstr Surg 130(1):53–63
23. Blanton MW et al (2009) Adipose stromal cells and platelet-rich
plasma therapies synergistically increase revascularization during
wound healing. Plast Reconstr Surg 123(2 Suppl):56S–64S
24. Lu F et al (2008) Improved viability of random pattern skin flaps
through the use of adipose-derived stem cells. Plast Reconstr
Surg 121(1):50–58
25. Natesan S et al (2011) Debrided skin as a source of autologous
stem cells for wound repair. Stem Cells 29(8):1219–1230
26. Natesan S et al (2011) A bilayer construct controls adipose-
derived stem cell differentiation into endothelial cells and peri-
cytes without growth factor stimulation. Tissue Eng Part A
17(7–8):941–953
27. Cao Y et al (2005) Human adipose tissue-derived stem cells
differentiate into endothelial cells in vitro and improve postnatal
neovascularization in vivo. Biochem Biophys Res Commun
332(2):370–379
28. Fischer LJ et al (2009) Endothelial differentiation of adipose-
derived stem cells: effects of endothelial cell growth supplement
and shear force. J Surg Res 152(1):157–166
29. da Silva Meirelles L, Caplan AI, Nardi NB (2008) In search of
the in vivo identity of mesenchymal stem cells. Stem Cells
26(9):2287–2299
30. Gimble JM, Katz AJ, Bunnell BA (2007) Adipose-derived stem
cells for regenerative medicine. Circ Res 100(9):1249–1260
31. Rehman J et al (2004) Secretion of angiogenic and antiapoptotic
factors by human adipose stromal cells. Circulation 109(10):
1292–1298
32. Rubina K et al (2009) Adipose stromal cells stimulate angio-
genesis via promoting progenitor cell differentiation, secretion of
angiogenic factors, and enhancing vessel maturation. Tissue Eng
Part A 15(8):2039–2050
33. Song SY, Chung HM, Sung JH (2010) The pivotal role of VEGF
in adipose-derived-stem-cell-mediated regeneration. Expert Opin
Biol Ther 10(11):1529–1537
34. Brem H et al (2009) Mechanism of sustained release of vascular
endothelial growth factor in accelerating experimental diabetic
healing. J Invest Dermatol 129(9):2275–2287
35. Wise LM et al (2012) The vascular endothelial growth factor
(VEGF)-E encoded by orf virus regulates keratinocyte prolifer-
ation and migration and promotes epidermal regeneration. Cell
Microbiol 14(9):1376–1390
36. Kidoya H et al (2008) Spatial and temporal role of the apelin/APJ
system in the caliber size regulation of blood vessels during
angiogenesis. EMBO J 27(3):522–534
37. Chiu LL, Radisic M (2010) Scaffolds with covalently immobi-
lized VEGF and Angiopoietin-1 for vascularization of engineered
tissues. Biomaterials 31(2):226–241
38. Janmey PA, Winer JP, Weisel JW (2009) Fibrin gels and their
clinical and bioengineering applications. J R Soc Interface
6(30):1–10
39. van Hinsbergh VW, Collen A, Koolwijk P (2001) Role of fibrin
matrix in angiogenesis. Ann N Y Acad Sci 936:426–437
40. Lesman A et al (2011) Engineering vessel-like networks within
multicellular fibrin-based constructs. Biomaterials 32(31):7856–
7869
Angiogenesis (2013) 16:745–757 757
123