Differential Healing Activities of CD34ⴙ and CD14ⴙ
Endothelial Cell Progenitors
Ola Awad, Eduard I. Dedkov, Chunhua Jiao, Steven Bloomer, Robert J. Tomanek, Gina C. Schatteman
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Objective—Peripheral blood contains primitive (stem cell-like) and monocytic-like endothelial cell progenitors. Diabetes
apparently converts these primitive progenitors, from a pro-angiogenic to anti-angiogenic phenotype. Monocytic
progenitors seem to be less affected by diabetes, but potential pro-angiogenic activities of freshly isolated monocytic
progenitors remain unexplored. We compared the ability of primitive and monocytic endothelial cell progenitors to
stimulate vascular growth and healing in diabetes and investigated potential molecular mechanisms through which the
cells mediate their in vivo effects.
Methods and Results—Human CD34⫹ primitive progenitors and CD14⫹ monocytic progenitors were injected locally into
the ischemic limbs of diabetic mice. CD14⫹ cell therapy improved healing and vessel growth, although not as rapidly
or effectively as CD34⫹ cell treatment. Western blot analysis revealed that cell therapy modulated expression of
molecules in the VEGF, MCP-1, and angiopoietin pathways.
Conclusions—Injection of freshly isolated circulating CD14⫹ cells improves healing and vascular growth indicating their
potential for use in acute clinical settings. Importantly, CD14⫹ cells could provide a therapeutic option for people with
diabetes, the function of whose CD34⫹ cells may be compromised. At least some progenitor-induced healing probably
is mediated through increased sensitivity to VEGF and increases in MCP-1, and possibly modulation of angiopoietins.
(Arterioscler Thromb Vasc Biol. 2006;26:758-764.)
Key Words: angiogenesis 䡲 CD34 䡲 diabetes 䡲 endothelial progenitor cells 䡲 monocytes
that CD14⫹ and CD14⫺ PBMCs have similar abilities to
differentiate into EC in vitro, although the CD14⫺ population
contained CD34⫹ PBMCs.11
Surprisingly, little is known about the effect of freshly
isolated exogenous monocytic progenitors on revascularization in vivo. Locally injected freshly isolated CD34⫺ PBMCs,
which contain ⬇10% monocytes, do not improve flow
restoration in ischemic limbs of nondiabetic and diabetic
mice. However, because in vitro studies demonstrate that
CD34⫺CD14⫺ cells inhibit CD14⫹ cell responsiveness, this is
not unexpected.18 Intravenous injection of freshly isolated
CD14⫹, CD34⫺, or total PBMCs in the nondiabetic mouse
also has no effect on healing, but because many thousand fold
more of the analogous cells are already present in the mouse
blood stream, this should not be unexpected.11 In contrast,
intravenous injection of cultured CD14⫹, CD34⫺, or total
PBMCs dramatically improved flow in the limbs relative to
untreated controls, as did macrophages derived from CD14⫹
PBMCs, albeit to a lesser degree.11 These data suggest that
monocytic progenitors require priming to differentiate into
EC and/or promote vascular growth. This should not be
surprising because monocytes require activation to perform
P
eripheral blood contains primitive (stem cell-like) and
monocytic endothelial cell (EC) progenitors that can
differentiate into ECs in vitro and integrate into the vasculature in vivo. However, in most situations, integration of
locally injected EC progenitors is rare. Nevertheless, EC
progenitors are of great clinical interest because they have
been shown to promote vascular growth and tissue healing.
Among primitive EC progenitors are human CD34⫹ peripheral blood mononuclear cells (PBMCs) and mouse bone
marrow Sca-1⫹lin⫺ cells.1–5 These cells leave the bone marrow and enter the circulation at a very low rate, representing
⬍0.1% of circulating cells. When injected locally, the freshly
isolated cells are potent stimulators of vessel growth in
ischemic tissue, although they rarely integrate into vessels.1–7
Human monocytes also differentiate into ECs in vitro8 –13
and in vivo,9,11,14,15 and CD14⫹ monocytes may be the
primary source of EC progenitors in the circulation.9 Moreover, a recent study concluded that monocytes are the
predominant source of “EPCs,”12 that is, bone marrowderived mononuclear cells that attach to fibronectin, bind
Ulex lectin, and take up acetylated low-density lipoprotein
after several days in culture.16,17 However, one study found
Original received June 21, 2005; final version accepted January 4, 2006.
From the Departments of Anatomy and Cell Biology (O.A., E.D., R.J.T.) and Exercise Science (C.J., S.B., G.C.S.), University of Iowa, Iowa City, Iowa.
Consulting Editor for this article was Alan M. Fogelman, MD, Professor of Medicine and Executive Chair, Departments of Medicine and Cardiology,
UCLA School of Medicine, Los Angeles, Calif.
O.A. and E.D. contributed equally to this work.
Correspondence to Gina Schatteman, PhD, Exercise Science FH412, University of Iowa, Iowa City, IA 52242. E-mail [email protected]
© 2006 American Heart Association, Inc.
Arterioscler Thromb Vasc Biol. is available at http://www.atvbaha.org
758
DOI: 10.1161/01.ATV.0000203513.29227.6f
Awad et al
CD14ⴙ Endothelial Cell Progenitors Promote Healing
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virtually every function with which they are associated.
Apparently, tissue culture can provide such a stimulus.
From a therapeutic standpoint, the need to culture cells
before use is not ideal. However, earlier studies with freshly
isolated human CD34⫹ and mouse Sca-1⫹ cells indicate that
when injected directly into the tissue, primitive EC progenitors require no priming, or if they do, the ischemic tissue
provides the signal. The same might be true for CD14⫹ cells.
Diabetes induces endothelial cell progenitor dysfunction,
including in CD34⫹ cells.1,4,5,19 –21. We examined the ability
of freshly isolated CD14⫹ cells to accelerate healing because
monocytic progenitors appear to be less negatively affected
by this disease than primitive progenitors.4 Under basal
conditions mouse diabetic and nondiabetic primitive progenitors were indistinguishable with respect to their ability to
proliferate and produce differentiated progeny. However,
when subjected to oxidative stress or hypoxia, the primitive
diabetic progenitors, but not their monocytic progeny, exhibited a reduced ability to produce endothelial cells relative to
their nondiabetic counterparts. Additional reasons for studying CD14⫹ cells are that they are abundant in the blood and
would be easy to harvest clinically, and in vitro data suggest
that they may be more potent stimulators of vascular growth
than whole PBMCs.18 Because CD34⫹ and CD14⫹ EC
progenitors have distinct in vitro characteristics, we also
tested whether the cells might cooperate to promote vascular
growth and healing in vivo.
The data suggest that CD14⫹ PBMCs may offer a therapeutic alternative for people with diabetes, the function of
whose CD34⫹ PBMCs is compromised. Intramuscularly injected freshly isolated CD14⫹ and CD34⫹ EC progenitors, or
the combination of the 2 increased aggregate arteriolar
density and promoted muscle salvage in the diabetic mouse
ischemic hindlimb. All cell treatments also accelerated blood
flow restoration, but with different kinetics. Western blot
analysis showed distinct patterns of pro-angiogenic factor
expression in CD34⫹ and CD14⫹ cell-treated limbs, which
may account for the different kinetics in blood flow
restoration.
Materials and Methods
Detailed procedures can be found in the online data supplement
(http://atvb.ahajournals.org). P⬍0.05 was considered statistically
significant for all assays.
Isolation and Labeling of PBMCS
Blood was collected from healthy adult volunteers according to
University of Iowa Institutional Review Board approved protocols
with informed consent. Cells were isolated as described.1 Briefly,
mononuclear cells were collected by density centrifugation. CD34⫹
PBMCs were isolated using 2 rounds of anti-CD34⫹ cell bead
selection. Residual CD34⫺ cells were used directly, or CD34⫺CD14⫹
PBMCs (hereafter referred to as CD14⫹ PBMCs for clarity) were
isolated by 2 rounds of anti-CD14⫹ cell bead selection. Some cells
were labeled with CM-DiI as described.9 Additional cells were
analyzed for expression of the vascular endothelial cell growth factor
receptor 2 (VEGFR2) (Figure I, available online at
http://atvb.ahajournals.org)
Animals
Animal procedures were approved by the University of Iowa Animal
Care and Use Committee. Anesthesia was induced with 4% isoflu-
759
Figure 1. LASER Doppler analysis of blood flow in ischemic diabetic mouse hindlimbs injected with PBMCs or vehicle on the
day of femoral artery ligation. Data expressed as percent flux in
ischemic limb relative to contralateral control limb. A, Approximate site of ligation (*) and PBMC injection (X). Outer tracing
indicates typical region of the limbs analyzed for blood flow.
Inner tracing approximates area of harvested muscle. B, Flow
after injection with vehicle (n⫽7), CD14⫹ PBMCs (CD14⫹) (n⫽5),
or a combination of CD14⫹ and CD34⫹ PBMCs (CD14⫹/CD34⫹)
(n⫽5). Both CD14⫹ injected groups differ from vehicle controls
beginning at day 8 (P⬍0.05). C, Flow in ischemic limbs after
vehicle or CD34⫺ PBMC injection. D, Effects of CD14⫹ PBMCs
(either singly or in combination) (n⫽10) on flow are intermediate
between those of control (CD34⫺ and vehicle) (n⫽11) and
CD34⫹ PBMC-injected mice (n⫽8). Flow in both the CD34⫹ and
CD14⫹ PBMC-injected groups differ from controls (P⬍0.01 and
P⬍0.05, respectively). P⫽0.075 for CD34⫹ versus CD14⫹ PBMC
treatment. Error bars⫽SEM.
rane and maintained with 0.8% to 1.2% isoflurane. Euthanasia was
performed by injection of 150 mg/kg sodium pentobarbital.
Diabetes was induced with streptozotocin in 8 to 12 weeks old
HFh11nu athymic mice male mice as described.22 Three to 4 weeks
after inducing diabetes, the left proximal femoral artery was ligated
as described.1 Two to 4 hours later, fresh human CD34⫹ PBMCs
(5⫻105, n⫽12 or 1⫻106; n⫽3), CD14⫹ PBMCs (1⫻106, n⫽11), or
the combination of the 2 (5⫻105 CD34⫹ and 1⫻106 CD14⫹; n⫽9)
were injected intramuscularly into the ischemic limbs. (Figure 1A)
Additional mice were injected with CD34- PBMCs (1⫻106; n⫽4),
with vehicle (n⫽7), or nothing (n⫽5). We saw no additional
acceleration of or increase in blood flow restoration when 1⫻106
rather than 5⫻105 CD34⫹cells were used. Thus, we did all subsequent experiments at the lower dose.
Blood Flow Analysis
Limb blood flow restoration distal to the ligation was analyzed
immediately before and after surgery and 2, 4, 6, 8, 10, and 12 days
after surgery using scanning LASER Doppler analysis as described1,23 (Figure 1A). Flow comparisons over time among groups
were performed by repeated measures ANOVA with Tukey’s posthoc analysis. Data are presented as percent mean blood flux in the
operated limb relative the unoperated limb.
Histology, Immunolabeling, and Morphometry
Control and ischemic limb hamstring muscles, from ⬇4 mm proximal to 4 mm distal of the injection site (Figure 1A) were collected 5
days after surgery (n⫽4 each group). Sections from 6 different
equally spaced levels were used for morphological examination and
arteriolar density measurements. Hematoxylin and eosin stained
760
Arterioscler Thromb Vasc Biol.
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sections were scored for inflammation,24 and the area of necrotic and
viable muscle measured. Data were analyzed by ANOVA with SNK
post-hoc analysis. The presence of mouse monocyte/macrophages
was assessed by incubation with anti-MMG lectin (Figure II,
available online at http://atvb.ahajournals.org).
To analyze intramuscular arteriolar parameters, smooth muscle
cells were labeled with anti-smooth muscle actin and the borders of
muscle fibers were delineated using rabbit anti-laminin antibodies.
Measurements were made on digitized images of 3 to 5 crosssections of the entire hamstring muscle group per animal in each
treatment condition using a modification of a previously described
method.25 Numerical density (vessels per area), vessel diameter
(external diameter), and vessels per muscle fiber were determined.
The axial ratios of vessels were used to calculate length density
according to the relationship: Length density⫽(N/A) ⫻ (⌺a/b)/N.
Where N is the number of vessels; a and b are the long and short
axes, respectively; and A is the area of the muscle. Aggregate length
density was calculated as the product of percent healthy muscle and
length density. Data were analyzed by ANOVA followed by Tukey’s
post-hoc test. These sections also were examined for the presence of
CM-DiI labeled cells.
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Western Blots
Hamstring muscles (Figure 1A) from uninjected or PBMC treated
ischemic limbs were collected 1 or 5 days after surgery; 50 ␮g
protein aliquots were separated by SDS-PAGE. After transfer,
membranes were probed with anti- angiopoietin-1, angiopoietin-2,
tie-2, vascular endothelial growth factor-A (VEGF-A), flk-1 (mouse
VEGFR2), or monocyte chemoattractant protein-1 (MCP-1) antibodies. Data were normalized to GAPDH levels. Samples from 4 to 6
mice in each group were analyzed 2 to 4 times each. Means were
compared by ANOVA with Tukey’s post-hoc analysis.
Results
ⴙ
CD14 PBMCs Accelerate Blood Flow Restoration
Bone marrow and circulating cells have distinct in vivo
characteristics.15 Because circulating cells would actually
enter sites of injury, and they are the ones most easily
harvested clinically, PBMCs were studied. Ischemic limbs of
diabetic mice were injected with vehicle or a subset of
PBMCs, hindlimb blood flow followed for 12 days, and the
percentage of flow restored calculated1 (Figure 1).Mean
blood glucose levels in diabetic mice were 435⫾35 mg/dL
(range 257 to 560). Nondiabetic mice of the same strain
averaged 115⫾6 mg/dL (range, 94 to 135; n⫽9).
In the first 6 days, there were no significant differences in
flow in limbs treated with CD14⫹ PBMCs or the CD14⫹/
CD34⫹ combination compared with vehicle controls, but
there was a trend toward increased flow (Figure 1B). By 8
days, both cell-treated groups showed increased flow relative
to vehicle treated controls (P⬍0.05), and this difference was
maintained through 12 days for CD14⫹ cell-treated limbs. At
no time did treatment with CD14⫹ PBMCs or the CD14⫹/
CD34⫹ PBMC combination differ from one another.
Though some data suggest otherwise,9,12 it has been reported that CD14⫹ and CD14⫺ PBMCs have similar abilities
to differentiate into EC in vitro.11 Thus, CD14⫹ and CD34⫺
PBMCs, which contain ⬇10% CD14⫹ and ⬇90% CD14⫺
PBMCs, could be equally potent in restoring blood flow.
However, consistent with our previous work, injection of
CD34⫺ PBMCs did not improve limb blood flow in these
mice (Figure 1C).
Flow restoration in limbs of mice that received CD14⫹
PBMCs either alone or in combination with CD34⫹ PBMCs,
Inflammatory Assessment of Ischemic Muscle
Treatment
Cellular infiltration/
inflammation*
P (vs vehicle)
⫹
Vehicle
CD14
CD14⫹/CD34⫹
CD34⫹
1.25⫾0.2
1.5⫾0.5
2.5⫾0.3
1.2⫾0.2
0.96
0.03
1.0
*Scale of 0 to 3 (no to extensive inflammatory infiltrate).
Mean⫾SEM; n⫽4 per group.
was intermediate between that of the poorly recovering
controls and rapidly healing CD34⫹ PBMC-treated mice
(Figure 1D). Also, CD14⫹ PBMCs did not induce significant
flow enhancement until day 8, whereas flow improved by day
2 in CD34⫹ PBMCs-treated limbs. Nevertheless, at 12 days
after treatment, mice receiving CD14⫹ PBMCs alone or in
combination with CD34⫹ cells exhibited a mean blood flow
restoration of 62% compared with 48% in controls, a 30%
improvement in resting flow (Figure 1D). Thus, freshly
isolated CD14⫹ cells potentiate blood flow restoration in
diabetic mice.
CD34ⴙ and CD14ⴙ PBMCs Promote
Muscle Healing
To determine whether blood flow restoration correlates with
increased muscle salvage, we examined hematoxylin and
eosin stained sections from 5 equally spaced regions for
inflammation, muscle necrosis, and muscle repair.24 Differences in inflammatory cell infiltration were not noticeably
different among groups, except in those co-injected with
CD14⫹ and CD34⫹ PBMCs wherein inflammation was extensive, even in regions where muscle was healthy (Table;
Figure 2D).
Vehicle-treated limbs exhibited marked necrosis. In some
areas, large segments of muscle were missing, resulting in a
small muscle mass. Regenerating muscle fibers with centralized nuclei were rare in these limbs. In contrast, minimal
necrosis, fibrosis, and inflammation were seen in limbs
treated with CD34⫹ PBMCs. In many regions these limbs
were indistinguishable from nonischemic controls (Figure 2A
and 2B). Even when single muscles were examined at
multiple levels, typically, the muscle was healthy with no
signs of necrosis (Figure 3A). Moreover, when entire crosssections of the harvested muscle group were examined, only
small localized foci of necrosis were apparent in the limbs of
all but one animal (Figure 3B). Necrosis and more extensive
fibrosis were apparent in limbs treated with CD14⫹ PBMCs
and in CD14⫹/CD34⫹ PBMC co-injected limbs (Figure 2C
and 2D). The only notable difference between CD14⫹ PBMC
and CD14⫹/CD34⫹ PBMC-treated limbs was the extensive
inflammation in the latter (Figure 2D). This appeared to be
caused by recruitment of endogenous cells, because mouse
monocytes, but few CM-DiI labeled cells, were present at
sites of inflammation (Figure II). Muscles with regenerating
fibers, characterized by central nuclei, were observed in all
cell treated groups (Figure 2E). CM-DiI labeled cells that
appeared to be integrated into capillaries could be seen, but
they were rare, consistent with previous reports of humanderived bone marrow cells in mouse vasculature.6,7,9
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Figure 2. Mouse muscle histology. Brightfield micrographs of 7 ␮m hematoxylin and eosin stained sections of hamstrings harvested 5
days after induction of ischemia. A, Muscle in uninjured limb. B, Morphologically healthy muscle in CD34⫹ PBMC-treated ischemic
limb. C, CD14⫹ PBMC-treated ischemic limb with necrosis and fibrosis (arrow). D, CD14⫹/CD34⫹ PBMC co-injected muscle in region of
extensive inflammatory infiltration and necrosis. E, Regenerating muscle fibers with centralized nuclei (arrowheads) in CD14⫹/CD34⫹
PBMC co-injected muscle. Inset shows regenerated muscle fibers with centralized nuclei (arrowheads) in cross-section. Bar is 100 ␮m
for all panels except for inset bar is 50 ␮m. F, Area of necrotic muscle as percentage of total muscle area in uninjected control (Cont)
and CD34⫹, CD14⫹, and CD34⫹/CD14⫹ cell-injected ischemic muscle 5 days after surgery. Bars indicate SEM. P⬍0.01 solid bracket.
P⬍0.05 dashed bracket. N⫽4 per group.
To quantitate histological findings, the areas of necrotic
and viable muscle were measured and the percentage of
necrotic muscle calculated. All cell treatments significantly
decreased necrosis, but CD34⫹ PBMCs treatment tended to
have the greatest effect (Figure 2F). Nevertheless, freshly
isolated CD14⫹ cells preserved muscle.
PBMC Therapy Modulates
Proangiogenic Pathways
changes 1 day after induction of ischemia (Figure 5A).
Protein levels in CD14⫹ cell-treated muscle were measured
for comparison. VEGF-A protein increased 2.5-fold in response to CD34⫹ PBMCs, but CD14⫹ PBMCs had no effect
on levels of this molecule. Both cell types profoundly
increased VEGFR2 protein. Whereas CD14⫹ cells elevated
ang-1 levels by 1.7-fold, CD34⫹ cells had no effect on either
ang-1 or ang-2, and neither cell modulated Tie-2 (the angiopoietin receptor) protein levels. Thus, both CD14⫹ and
CD34⫹ PBMC treatment profoundly increased sensitivity to
VEGF family ligands, but only CD14⫹ cells affected the
angiopoietin pathway.
CD14⫹ cell and CD34⫹/CD14⫹ cell flow-mediated changes
become apparent several days after injection, so we measured
angiogenic protein levels in ischemic limbs five days after
treatment (Figure 5B). As at day 1, VEGF-A levels were
similar in untreated and CD14⫹ PBMC-treated muscle,
whereas VEGF-A was higher than controls in CD34⫹/CD14⫹
and CD34⫹ cell-treated muscle. However, VEGFR2 protein
declined to undetectable levels in all groups by this time.
Angiopoietin levels varied widely among individual animals
at day 5, whereas Tie 2 levels were consistent across animals,
and tended to decrease in cell-injected muscle. Only in
CD34⫹/CD14⫹ cell co-injected mice was this decrease statistically significant.
MCP-1 has pleiotropic effects on EC progenitors and
vascular growth and remodeling, so we also measured relative levels of MCP-1. Cell treatment had no effect on MCP-1
levels 1 day after surgery, but by 5 days the level of MCP-1
protein was increased 2.4-, 6.0-, and 14.8-fold relative to
controls in CD14⫹, CD34⫹/CD14⫹, and CD34⫹ cell-treated
muscle, respectively (Figure 5C).
Cell injection-induced changes in protein levels in the VEGF
and angiopoietin (ang) systems were measured in the ischemic muscle. Because CD34⫹ PBMC therapy rapidly induces
increases in blood flow, we measured CD34⫹ cell mediated
We compared direct local injection of freshly isolated CD14⫹
and CD34⫹ PBMCS, and found that while CD14⫹ monocytic
PBMC Treatment Increases Aggregate
Vascular Density
LASER Doppler analysis assesses collateral formation, but
muscle salvage also requires an adequate microvasculature.
Hence, we measured arteriolar parameters in cross-sections of
the hamstring muscles at three similar levels in viable muscle
in all groups. Capillary density is often used to probe the
microvasculature, but capillaries are notoriously unstable in
healing tissue. Further, arteriolar density gives insight into the
pool of vessels available for remodeling into collaterals.
No increase in vessel diameter as a result of ischemia or
cell treatment was observed (Figure 4A). There was a trend
(P⫽0.08) toward an ischemia-induced increase in the vessel
to fiber ratio, but again, there was no significant effect of cell
treatment (Figure 4B). Ischemia was associated with increased arteriolar numerical and length density in viable
muscle, but neither of these parameters was affected by cell
treatment (Figure 4C and 4D). However, necrotic muscle was
essentially devoid of arterioles in all treatment groups.
Because of this, cell treatment resulted in greater aggregate
arteriolar densities than in untreated limbs, because there was
more viable (ie, less necrotic) muscle (Figure 4E; Figure III,
available online at http://atvb.ahajournals.org).
Discussion
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Figure 3. CD34⫹ PBMC-treated ischemic muscle. Brightfield
micrographs of 7 ␮m hematoxylin and eosin stained hamstrings
sections harvested 5 days after induction of ischemia. A, Series
of sections from a single muscle region at ⬇100-␮m intervals.
Note lack of necrotic tissue. Some inflammation can be seen by
the presence of nuclei (black dots) intercolated between the
muscle fibers. Bar⫽25 ␮m. B, Composite micrograph of a
cross-section of the entire hamstring group. Note healthy
appearance of the muscle except in small region (inset) where
necrosis and regeneration are present. Bar⫽1 mm.
EC progenitors promote healing, accelerate blood flow restoration, and induce vascular growth, they do so less well
than their CD34⫹ primitive EC progenitor counterparts.
Nevertheless, because diabetes appears to have more deleterious effects on primitive than monocytic progenitor function,
CD14⫹ PBMCS may provide an alternative therapeutic tool
for this large subset of patients. Moreover, CD14⫹ therapy
may be feasible in an acute setting, because CD14⫹ PBMCs
are abundant and easy to isolate, and require no pre-activation
when directly injected into the affected tissue.
Hematopoietic stem cells and monocytes both promote
vascular growth.26,27 Thus, we expected that co-injection of
CD34⫹ and CD14⫹ PBMCs would lead to the greatest muscle
salvage and tissue vascularization. The data do not show this
(Figure 1), and our histological analysis suggests that excessive inflammation (Table) may be the reason. The cellular
infiltrate probably reduces the sensitivity of the Doppler flow
measurements, leading to an underestimate of true blood
flow. More importantly, as the vascular density and muscle
salvage data imply, although inflammation can be an angiogenic stimulus, excessive inflammation may have damped the
limbs’ ability to make new or remodel existing vessels.
Interestingly, the inflammation does not appear to be caused
by the presence of the human cells per se, because we found
few human cells in the tissue at 5 days, when inflammation
was extensive. Rather, it seems that the exogenous cells act in
concert to attract endogenous cells, possibly through MCP-1.
The Doppler blood flow measurements indicate that cell
therapy increases flow to collaterals (Figure 1). This could be
caused in part by vasodilation initially, but as the blood flow
remains elevated 12 days after induction of ischemia (by
which time inflammation is largely absent), remodeling
appears to occur ultimately. Arteriolar density and size also
influence intramuscular limb blood flow, but these have not
been examined previously in PBMC treated limbs. Ischemia
induces an increase in arteriolar density but not size, and,
surprisingly, cell therapy does not further increase arteriolar
density or alter size in viable muscle (Figure 4). Apparently,
once a critical arteriolar density necessary to preserve ischemic muscle is reached, additional vessels are not assembled.
However, because there is more viable muscle in PBMC than
vehicle-treated limbs, PBMC treatment results in an increased
aggregate vessel density (Figure 2F; Figure III). Thus, rapid
and extensive vascular growth must occur in the PBMCtreated limbs.
CD34⫹ and CD14⫹ PBMCs might exert some of their
effects through modulation of pro-angiogenic molecules, and
CD34⫹ but not CD14⫹ PBMCs increase VEGF-A protein
(Figure 5A). We do not know if the VEGF detected was
produced by the exogenous or endogenous cells. However,
VEGF is a highly conserved protein across species, and
human VEGF cross-reacts in the mouse.28 Whether produced
by the injected human or by mouse cells the protein should be
Figure 4. Vascular parameters in hamstring
muscles. Measurements of smooth muscle
actin positive vessels in uninjured muscle
and viable regions of hamstring muscles 5
days after surgery in control and CD34⫹,
CD14⫹, and CD34⫹/CD14⫹ cell-treated
limbs. A, Mean radius of vessels. B, Vessels
per 100 muscle fibers (ANOVA P⫽0.08). C,
Number of vessel per cross-sectional viable
muscle area. D, Vessel length per volume of
viable muscle. E, Vessel density in all
(necrotic and viable) muscle expressed as
fold relative to untreated controls. Bars indicate SEM. N⫽3 to 4 per group. *P⬍0.05
relative to all other groups.
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Figure 5. Relative protein levels of
angiogenic factors and their receptors in
ischemic limbs. Fold change in protein
level in ischemic muscle in CD34⫹,
CD14⫹, and CD34⫹/CD14⫹ PBMCtreated relative to untreated ischemic
control (Cont) muscle. Changes in VEGF
and the angiopoietin system (A) 1 day
and (B) 5 days after surgery and cell
treatment. C, Changes in MCP-1 protein
1 day (left) and 5 days (right) after surgery and cell treatment. Bars indicate
SEM. P⬍0.01 solid bracket. P⬍0.05
dashed bracket. N⫽4 to 6 mice per
group.
active in vivo in the mouse. More significantly, both cells
augment responsiveness to VEGF family proteins by dramatically increasing VEGFR2 protein. The potent induction of
VEGF-A and VEGFR2 may contribute to the strong early
vascular growth response to CD34⫹ PBMCs compared with
CD14⫹ PBMCs.
Although VEGF-A levels remained slightly elevated in
limbs treated with CD34⫹ cells at day 5, we were unable to
detect VEGFR2 protein (Figure 5B). This is not a technical
problem as the same tissue samples were used for analyses of
other proteins, and VEGFR2 was detected in day 1 samples.
The levels simply had declined, suggesting that most new
vessel growth is completed by this time, and that subsequent
improvements in flow are caused by remodeling. This may
explain why the greatest increases in limb blood flow occur
within the first 6 days after induction of ischemia. This too
indicates that for acute events (such as myocardial infarction)
PBMC-based therapies may be most effective when used
soon after the event, emphasizing the importance of being
able to use freshly isolated cells therapeutically.
The angiopoietin data are more complex. CD34⫹ cells have
no apparent immediate effect on the angiopoietins or their
receptor, although CD14⫹ cells induce an increase in angiopoietin 2, suggesting that these cells may have an early
influence on remodeling (Figure 5A). The day 5 data were
striking because, although tie-2 receptor levels were consistent from animal to animal and cell therapy tended to depress
receptor levels, angiopoietin 1 and 2 protein levels varied
widely among cell-treated mice (Figure 5B). Protein levels in
individual mice were either similar to controls, or in the case
of angiopoietin 1, much lower, or angiopoietin 2, much
higher. Thus, angiopoietin modulation seems to be dynamic
at this point in time, consistent with vascular remodeling.
MCP-1 recruits monocytes, induces EC progenitor differentiation, and promotes arteriogenesis.14,15,29,30 Hence, the
dramatic elevation in MCP-1 at day 5, but not at day 1, may
further explain why improvements in blood flow are not
apparent in CD14⫹ treated mice until day 6 (Figure 5C). That
is, late elevation of MCP-1 may contribute to the increased
arteriogenesis. Additionally, MCP-1 could be recruiting endogenous EC progenitors and inducing their differentiation,
thereby increasing microvascular growth.
In sum, our data indicate that local injection of small
numbers of freshly isolated circulating cells can dramatically
impact healing and vascular growth, but that CD34⫹ PBMCs
are more effective than CD14⫹ cells in doing so. Nevertheless, CD14⫹ cells could offer a therapeutic alternative for
people with diabetes, the function of whose CD34⫹ PBMCs
may be compromised.1,31 Finally, the data suggest that at least
some PBMC induced healing is mediated through the VEGF
and MCP-1 pathways, and possibly the angiopoietins.
Acknowledgments
This work was supported by grants from the Juvenile Diabetes
Research Foundation 1-2001 to 534 (G.S.), National Institutes of
764
Arterioscler Thromb Vasc Biol.
April 2006
Health DK55965 (G.S.), and HL075446 (R.T.). The authors thank
Sonny Patel for technical assistance.
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9. Harraz M, Jiao C, Hanlon HD, Hartley RS, Schatteman GC. Cd34(-)
blood-derived human endothelial cell progenitors. Stem Cells. 2001;19:
304 –312.
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Strasser RH, Daniel WG. Monocytes coexpress endothelial and macrophagocytic lineage markers and form cord-like structures in Matrigel under
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11. Urbich C, Heeschen C, Aicher A, Dernbach E, Zeiher AM, Dimmeler S.
Relevance of monocytic features for neovascularization capacity of circulating endothelial progenitor cells. Circulation. 2003;108:2511–2516.
12. Rehman J, Li J, Orschell CM, March KL. Peripheral blood “endothelial
progenitor cells” are derived from monocyte/macrophages and secrete
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13. Zhao Y, Glesne D, Huberman E. A human peripheral blood monocytederived subset acts as pluripotent stem cells. Proc Natl Acad Sci U S A.
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14. Moldovan NI, Goldschmidt-Clermont PJ, Parker-Thornburg J, Shapiro
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Differential Healing Activities of CD34+ and CD14+ Endothelial Cell Progenitors
Ola Awad, Eduard I. Dedkov, Chunhua Jiao, Steven Bloomer, Robert J. Tomanek and Gina C.
Schatteman
Arterioscler Thromb Vasc Biol. 2006;26:758-764; originally published online January 12, 2006;
doi: 10.1161/01.ATV.0000203513.29227.6f
Arteriosclerosis, Thrombosis, and Vascular Biology is published by the American Heart Association, 7272
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Detailed Methods
Isolation, Labeling, and FACS of PBMCS
50-100 ml human blood was collected from healthy adult volunteers according to University of Iowa
Institutional Review Board approved protocols with informed consent as described.1 Briefly,
mononuclear cells were collected after fractionation on Histopaque 1077 (Sigma, St. Louis) per
manufacturer's instructions and washed. CD34+ PBMCs were isolated using 2 rounds of CD34+ cell
selection with an auto MACS (Miltenyi, Auburn, CA) per manufacturer’s instructions. Residual CD34cells were used directly or CD34-CD14+ PBMCs (hereafter referred to as CD14+ PBMCs for clarity) were
isolated by two rounds of CD14+ selection using auto MACS according to manufacturer’s instructions.
Prior to injection, cells were labeled with the vital dye, CM-DiI (Molecular Probes, Eugene, OR), as
described in some animals.2
To assess VEGFR2 expression on the freshly isolated CD14+ and CD34+ PBMCs, cells were incubated 30
min on ice with 20μg/ml anti-VEGFR2 (Abcam Cat # Ab9539, Cambridge, MA), washed twice in 1%
fetal bovine serum (FBS) in PBS then overlaid on FBS serum and pelleted. Cells were resuspended in
1% FBS in PBS and incubated 30 min with 13 μg/ml Alexa 488 anti mouse IgG then washed as above,
6
and resuspended in propidium iodide containing buffer at 1 x10 cells/ml in PBS for FACS.
Animals
Animal procedures were approved by the University of Iowa Animal Care and Use Committee.
Anesthesia was induced with 4% isoflurane and maintained with 0.8-1.2% isoflurane. Euthanasia was
performed by injection of 150mg/kg sodium pentobarbital.
Induction of Ischemia and Cell Treatment
Diabetes was induced with streptozotocin in 8-12 wk old HFh11nu athymic mice male mice (Jackson
Laboratories, Bar Harbor, ME) as described.3 Mice that lost more than 20% of their body weight or
appeared ill were euthanized and excluded from analysis. Three to four weeks after inducing diabetes, the
left proximal femoral artery was ligated as described.1 Two to four hours later, fresh human CD34+
PBMCs (5 x 105, n=12 or 1 x 106; n=3), CD14+ PBMCs (1 x 106, n=11), or the combination of the two (5
x 105 CD34+ and 1 x 106 CD14+; n=9) were injected intramuscularly into the ischemic limbs. (Figure 1A)
Additional mice were injected with CD34- PBMCs (1 x 106) (n=4), vehicle (n=7), or nothing (n=5).
Blood Flow Analysis
Limb blood flow restoration was assessed using scanning LASER Doppler analysis as described.1,4 Flow
was analyzed immediately before and after surgery and 2, 4, 6, 8, 10, and 12 days after surgery in the
entire limb distal to the ligation. (Figure 1A) Comparisons of blood flow over time among groups were
made by repeated measures ANOVA with Tukey’s honestly significant difference post-hoc analysis using
SPSS software (SPSS Science, Chicago, IL). P< 0.05 was considered statistically significant. Data are
presented as percent mean blood flux in the operated ischemic limb relative the unoperated control limb.
Histology, Immunolabeling, and Morphometry
Control and ischemic limb hamstrings muscles, from approximately 4mm proximal to 4mm distal of the
injection site (Figure 1A) were collected 5 days after surgery (n = 4 each group), methanol fixed, paraffin
embedded, and serially sectioned at 7 μm. Sections from 6 different equally spaced levels were used for
morphological examination and vessel density measurements. Six hematoxylin and eosin stained sections
were scored for inflammation,5 and the area of necrotic and non-necrotic muscle was estimated by tracing
using Metavue software (Universal Imaging, Downington, PA). Data were analyzed by ANOVA
followed by SNK post-hoc analysis.
To examine ischemic muscle for the presence of mouse monocyte/macrophages, additional sections of
muscle in which inflammation was present were incubated with 1μg/ml anti-mouse macrophage
galactose-specific lectin (MMGL) clone ER-MP23 (Abcam, Cambridge, MA) overnight followed by 2 h
with 10 μg/ml Alexa 488 anti-rat IgG (Molecular Probes, Eugene , OR).
To analyze vascularity, smooth muscle cells were labeled with Cy3-anti-smooth muscle actin (1:600;
Sigma, St. Louis, MO) 40 min at 37oC and the borders of muscle fibers were delineated by incubation
with rabbit anti-laminin (1:30; Sigma, St. Louis) 2 h at 37oC followed by Alexa 488 goat anti-rabbit
(Molecular Probes). Images were digitized with Image Pro software (Media Cybernetics, San Diego,
CA). Measurements were made on images of 3-5 cross-sections of the entire hamstrings muscle group
per animal in each treatment condition using a modification of a previously described method.6 The axial
ratios of vessels were used to calculate length density according to the relationship: Length density =
numerical density X a/b, where a and b are the long and short axes, respectively. Numerical density
(vessels per area), vessel diameter (external diameter), and vessels per muscle fiber were also determined.
Data were analyzed by ANOVA followed by Tukey’s honestly significant difference post-hoc test. These
sections were also examined for the presence of human CM-DI labeled cells.
Western Blot
Hamstrings muscle, (Figure 1A) from PBMC injected or uninjected ischemic limbs were collected one or
five days after induction of ischemia and snap frozen. Tissue was pulverized in liquid N2, resuspended in
lysis buffer (50mM Tris, pH 7.5, 1mM EDTA, 1% Triton, 0.9% NaCl, 1mM PMSF) clarified at 12,000 x
g for 30 min at 4oC. Clarified supernatants were used immediately or stored at -80oC until use. Protein
concentration were determined using a modified Bradford assay using Protein Assay Reagent (BioRad,
Hercules, CA).7 50 μg protein samples were separated by SDS-PAGE and transferred to PVDF (BioRad).
Membranes were blocked in 5% non-fat milk, incubated in primary antibody 2 hr at room temperature or
overnight at 4oC in blocking solution, washed 3 times in PBS with 0.05% /Tween-20, then incubated in
horseradish peroxidase conjugated secondary antibody for 1 hr at room temperature. Bands were
visualized with Super Signal (Pierce, Rockford, IL) after 3 washes in PBS with 0.05% /Tween-20.
Primary antibodies used were anti- angiopoietin-1 (1:300), angiopoietin-2 (1:200), tie-2 (1:200), VEGF
(1:2000), flk-1 (1:200), (all from Santa Cruz Biotechnology, Santa Cruz, CA), and MCP-1 (1:1500)
(eBioscience, San Diego, CA). Blots were analyzed using anti-GAPDH (1:8000) (Chemicon, Temecula,
CA) as above and data were normalized to GAPDH levels. Tissue from four to six mice in each group
was analyzed 2-4 times each. Means from each animal were compared among groups by ANOVA.
Tukey's post-hoc analysis was done with P< 0.05 considered statistically significant.
Figure I.
Figure I. Representative FACS plots from analysis of freshly isolated CD34+ and CD14+ PBMCs. A)
Anti-VEGFR2 reactivity was not detected on CD34+ PBMCs. B) Very low-level anti-VEGFR2 reactivity
was detected on 77.7 + 1.5% of CD14+ PBMCs in 3 analyses. The maximal level of fluorescence was
two orders of magnitude less than that found for HUVEC. (Data not shown.)
Figure II.
Figure II. Mouse muscle histology in areas of inflammation. Micrographs of 7 μm sections hamstrings
harvested five days post-induction of ischemia. Hematoxylin and eosin stained sections (A, C, E, & G)
and adjacent sections (B, D, F, & H) immunolabeled with anti-MMGL to identify mouse
monocyte/macrophages. A-B) Untreated, C-D) CD34+ treated, E-F) CD14+ PBMC treated, G-H) CD14+/
CD34+ PBMC co-injected limbs. Bar = 10μm.
Figure III.
Figure III. Schematic of aggregate arteriole density in PBMC (left) and vehicle (right) treated ischemic
muscles. The density of arterioles (black dots) is similar in viable muscle in both groups, but the amount
of viable muscle is greater in PBMC treated limbs than in vehicle controls. Aggregate arteriolar density
(arteriolar density x healthy muscle/all muscle) is greater in PBMC than vehicle treated limbs. Vehicle
treated limbs are small because of muscle loss due to necrosis.
References
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Schatteman GC, Hanlon HD, Jiao C, Dodds SG, Christy BA. Blood-derived angioblasts
accelerate blood-flow restoration in diabetic mice. J Clin Invest. 2000;106:571-578.
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Harraz M, Jiao C, Hanlon HD, Hartley RS, Schatteman GC. Cd34(-) blood-derived
human endothelial cell progenitors. Stem Cells. 2001;19:304-12.
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Kunjathoor VV, Wilson DL, LeBoeuf RC. Increased atherosclerosis in streptozotocininduced diabetic mice. J Clin Invest. 1996;97:1767-73.
4.
Couffinhal T, Silver M, Zheng LP, Kearney M, Witzenbichler B, Isner JM. Mouse model
of angiogenesis. Am J Pathol. 1998;152:1667-79.
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Akar H, Sarac A, Konuralp C, Yildiz L, Kolbakir F. Comparison of histopathologic
effects of carnitine and ascorbic acid on reperfusion injury. Eur J Cardiothorac Surg.
2001;19:500-6.
6.
Zheng W, Weiss RM, Wang X, Zhou R, Arlen AM, Lei L, Lazartigues E, Tomanek RJ.
DITPA stimulates arteriolar growth and modifies myocardial postinfarction remodeling.
Am J Physiol Heart Circ Physiol. 2004;286:H1994-2000.
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