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VASCULAR BIOLOGY
Tissue type plasminogen activator regulates myeloid-cell dependent
neoangiogenesis during tissue regeneration
*Makiko Ohki,1 *Yuichi Ohki,1 Makoto Ishihara,1 Chiemi Nishida,1 Yoshihiko Tashiro,1 Haruyo Akiyama,1
Hiromitsu Komiyama,1 Leif R. Lund,2 Atsumi Nitta,3 Kiyofumi Yamada,3 Zhenping Zhu,4 Hideoki Ogawa,5 Hideo Yagita,6
Ko Okumura,5 Hiromitsu Nakauchi,1 Zena Werb,7 †Beate Heissig,1,5,8 and †Koichi Hattori1,5
1Center
for Stem Cell Biology and Regenerative Medicine, Institute of Medical Science, University of Tokyo, Tokyo, Japan; 2Department of Cellular and
Molecular Medicine, Faculty of Health Science, University of Copenhagen, Copenhagen, Denmark; 3Department of Neuropsychopharmacology, Nagoya
University Graduate School of Medicine, Nagoya, Japan; 4ImClone Systems, New York, NY; 5Atopy (Allergy) Research Center, Juntendo University School of
Medicine, Tokyo, Japan; 6Department of Immunology, Juntendo University School of Medicine, Tokyo, Japan; 7Department of Anatomy, University of California,
San Francisco; and 8Frontier Research Initiative, Institute of Medical Science at the University of Tokyo, Tokyo, Japan
Ischemia of the heart, brain, and limbs is
a leading cause of morbidity and mortality worldwide. Treatment with tissue type
plasminogen activator (tPA) can dissolve
blood clots and can ameliorate the clinical outcome in ischemic diseases. But
the underlying mechanism by which tPA
improves ischemic tissue regeneration is
not well understood. Bone marrow (BM)–
derived myeloid cells facilitate angiogenesis during tissue regeneration. Here, we
report that a serpin-resistant form of tPA
by activating the extracellular proteases
matrix metalloproteinase-9 and plasmin
expands the myeloid cell pool and mobilizes CD45ⴙCD11bⴙ proangiogenic, myeloid cells, a process dependent on vascular endothelial growth factor-A (VEGF-A)
and Kit ligand signaling. tPA improves the
incorporation of CD11bⴙ cells into ischemic tissues and increases expression of
neoangiogenesis-related genes, including VEGF-A. Remarkably, transplantation
of BM-derived tPA-mobilized CD11bⴙ cells
and VEGFR-1ⴙ cells, but not carriermobilized cells or CD11bⴚ cells, acceler-
ates neovascularization and ischemic tissue regeneration. Inhibition of VEGF
signaling suppresses tPA-induced neovascularization in a model of hind limb
ischemia. Thus, tPA mobilizes CD11bⴙ
cells from the BM and increases systemic
and local (cellular) VEGF-A, which can
locally promote angiogenesis during ischemic recovery. tPA might be useful to
induce therapeutic revascularization in
the growing field of regenerative medicine. (Blood. 2010;115(21):4302-4312)
Introduction
The fibrinolytic system includes a broad spectrum of proteolytic
enzymes with physiologic and pathophysiologic functions in
several processes such as hemostatic balance, tissue remodeling,
tumor invasion, reproduction, and angiogenesis.
The serine protease plasmin is responsible for the degradation of
fibrin into soluble degradation products (fibrinolysis). Plasmin is generated through cleavage of the proenzyme plasminogen (Plg) by the
urokinase plasminogen activator (uPA) or tissue-type plasminogen
activator (tPA). tPA consists of a kringle- and trypsin-like serine protease
domain.1 The activity of uPA and tPA is regulated by specific plasminogen activator inhibitors. In the absence of fibrin, tPA displays low
activity toward Plg.2 In the presence of fibrin this activity is 2 orders of
magnitude higher. The catalytic efficiency of tPA for activation of
cell-bound Plg is approximately 10-fold higher than that in
solution. Most cells bind Plg through its lysine binding sites
with a high capacity but a relatively low affinity.3 Plg receptors
such as the integrin ␣M␤2 play an important role in macrophage
motility.4 CD11b/CD18 cells adhere to fibrin, but tPA by its
ability to bind to CD11b, has been shown to induce local
fibrinolysis and to render adherent cells into migrating cells.5
tPA has been shown to have numerous biologic functions. For
example, within the central nervous system (reviewed by Melchor
and Strickland6) tPA is expressed by neurons and microglial cells
(resident macrophages of the brain and spinal cord), where it can
generate plasmin to degrade a variety of nonfibrin substrates (eg,
␤-amyloid), can act as a direct protease without Plg involvement
(eg, for the activation of latent platelet-derived growth factor-CC),
or can function as a nonproteolytic modulator (eg, of the N-methylD-aspartate receptors).
Besides their fibrinolytic activities, plasmin and Plg activators
are also implicated in tissue proliferation and cellular adhesion,
because they can proteolytically degrade the extracellular matrix
(ECM) and regulate the activation of both growth factors and
matrix metalloproteinases (MMPs; for review, see Zorio et al7).
PAs and plasmin generation in specific microenvironments in the
bone marrow (BM) may be one of the factors orchestrating
hematopoiesis.8,9 Plg activation promotes the release of Kit ligand
from BM stromal cells.9,10 Plasmin can cleave thrombopoietin, the
master cytokine of megakaryopoiesis and platelet production,
thereby decreasing its biologic activity.11 Plasmin can further
change the release or activation status of basic fibroblast growth
factor, transforming growth factor ␤, and interleukin-1 ␤, which
either directly or indirectly modulates myelopoiesis by releasing
growth factors such as granulocyte colony-stimulating factor
Submitted August 3, 2009; accepted January 10, 2010. Prepublished online as
Blood First Edition paper, January 28, 2010; DOI 10.1182/blood-2009-08-236851.
The online version of this article contains a data supplement.
*M.O. and Y.O. contributed equally to this study.
†B.H. and K.H. share senior authorship.
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The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 USC section 1734.
© 2010 by The American Society of Hematology
BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
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BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
(G-CSF) or granulocyte macrophage CSF (GM-CSF), and macrophage CSF.12,13 Thus, paradoxically, plasmin generation can have
both a permissive and an inhibitory effect on hematopoiesis.
A number of cell model studies have shown that MMP
activation, notably activation of stromelysin-I, MMP-3, and MMP-9,
can occur at the cell surface through the uPA/uPAR/Plg cascade for
plasmin generation.14 We found increased soluble uPA receptor
serum levels in samples from patients treated with G-CSF.15
Endothelial cell invasion is an essential event during angiogenesis, the formation of new blood vessels. This process involves the
degradation of the ECM, the basement membrane, and the interstitial stroma and is governed by the activation of MMPs.16 During
angiogenesis, endothelial cells stimulated by hypoxia and angiogenic factors start to proliferate and to migrate through the ECM.
Endothelial cells are the main source of tPA in the blood circulation. tPA is released after injury or by brain endothelial cells on
monocyte interaction.17 Recent studies emphasized a role for
BM-derived CD45⫹ myeloid hematopoietic cells at ongoing angiogenic sites.18-20 Myeloid cells may locally secrete angiogenic
factors or MMP-9, which may in turn increase vascular endothelial
growth factor (VEGF-A) bioavailability.21 The beneficial effect of
recombinant tPA on tissue regeneration in ischemic diseases has
been attributed to its ability to degrade fibrin. Here, we report that
tPA promotes neoangiogenesis during hind limb (HL) ischemia by
mobilizing angiopotent CD45⫹CD11b⫹ myeloid cells coexpressing VEGF receptor-1 (VEGFR-1), a process dependent on VEGF
signaling. These tPA-mobilized cells showed improved tissue
incorporation into the ischemic niche, most probably because of
their increased expression of chemokine receptors such as CXCR4
and CCR2, resulting in faster tissue regeneration. tPA not only
increased the absolute number of CD11b⫹ myeloid cells but also
enhanced their angiogenic performance, making them a good
cellular target for cell-based treatment of ischemic diseases.
Methods
Animals
Plg⫹/⫹ and Plg⫺/⫺ mice and MMP9⫹/⫹ and MMP9⫺/⫺ mice were used after
more than 10 back crosses onto a C57BL/6, or CD1 background,
respectively.10 C57BL/6 mice were purchased from SLC. We used tPA⫹/⫹
and tPA⫺/⫺ mice and C57BL/6 mice, expressing green fluorescent protein
(GFP) under a ␤-actin promoter. Animal studies were approved by the
Animal Review Board of Juntendo University, Tokyo.
Chimeric BM transplantation model
Plg⫹/⫹ or Plg⫺/⫺ animals were lethally irradiated (900 cGy) and 6 hours
later were transplanted with 2 ⫻ 107 BM-derived mononuclear cells from
either Plg⫹/⫹ or Plg⫺/⫺ animals.
Study design
Mutant tPA (Eizai; resuspended in 150 ␮L of 0.2% bovine serum albumin
[BSA]; Sigma Chemical Co) was administered (10 mg/kg body weight) to
mice with or without induction of HL ischemia by daily intraperitoneal
injections from day 0 to day 2. Control mice received 150 ␮L of 0.2% BSA
or in some experiments G-CSF (200 ␮g/kg body weight from day 0 to day
2; Kirin Pharma Company). Recombinant murine VEGF-A (PeproTech)
was injected intraperitoneally (100 ng/mouse, daily from day 0 to day 2).
C57BL/6 mice were coinjected with tranexamic acid (Daiichi-Sankyo)
and tPA intraperitoneally (day 0-2). C57BL/6 mice were treated with uPA
(1 mg/kg body weight; Mitsubishi Tanabe Pharma) intraperitoneally from
day 0 to day 2.
tPA-DRIVEN MYELOID CELL DRIVEN TISSUE REGENERATION
4303
C57BL/6 mice were treated with a catalytically inactivated tPA
(HTPA-ALA; Molecular Innovation) or rtPA (Eizai) at a concentration of
1.32 mg/kg from day 0 to day 2. C57BL/6 mice received daily injection of
tPA (day 0-3) and were coinjected with batroxobin (20 BU/kg body weight
intraperitoneally) from day ⫺3 until day 5. Peripheral blood was harvested.
Fluorescence-activated cell sorting
Peripheral mononuclear cells (PBMCs) were stained with the following
antibodies: CD45-Pacific Blue (clone 30-F11; BioLegend), CD11b-FITC or
APC (clone M1/70; PharMingen), F4/80-FITC (clone BM8; BioLegend),
CD184-FITC (clone 2B11/CXCR4; PharMingen), and VEGFR-1-PE (clone
MF-1; ImClone Systems). Cells were analyzed with the use of a BD
FACSAria (Becton Dickinson). CD11b cell populations were isolated,
staining with CD45 and CD11b antibodies.
Reverse transcription PCR analysis
Total RNA was extracted with the use of RNA Trizol (Invitrogen). cDNA
was generated from the following cell populations isolated from PBMCs of
tPA- or BSA-PBS (phosphate-buffered saline)–treated mice: CD11b⫹ and
CD11b⫺ cells. cDNA was amplified by polymerase chain reaction (PCR) with
the following specific forward and reverse primer pairs: for VEGF-A,
5⬘-AATGCTTTCTCCGCTCTGAA-3⬘ and 5⬘-CAGGCTGCTGTAACGATGAA-3⬘; for CXCR-4, 5⬘-CTTGACTGGCATAGTCGGCAATGGA-3⬘ and 5⬘-TGCTGGAATTGAAACACCACCATCC-3⬘; for CCR2,
5⬘-AGAGGTCTCGGTTGGGTTGT-3⬘ and 5⬘-ATCATAACGTTCTGGGCACC-3⬘; for GM-CSF, 5⬘-TAGCTGATAAGGGCCAGGAG-3⬘;
for neuropilin-1, 5⬘-ACCAAGGAGACCACTGGAAAG-3⬘ and 3⬘AAGAGAGGAAAAAAGGGGGCT-5⬘; for Tie-2, 5⬘-GGCTGATTCTTCGGAGATGT-3⬘ and 3⬘-GAAAGGCTTTTCCACCATC-5⬘; and for
the ␤-actin control, 5⬘-TAAAACGCAGCTCAGTAACAGTCCG-3⬘ and
5⬘-TGGAATCCTGTGGCATCCATGAAAC-3⬘.
Murine ischemic HL mode
Mice were anesthetized with ketamine, 80 mg/kg body weight, and
xylazine, 10 mg/kg body weight, given intraperitoneally for operative
ligation of 1 femoral artery. Starting from the day of femoral ligation tPA
was injected intraperitoneally daily for a period of 3 days. Control groups
were similarly injected with 150 ␮L of 0.2% BSA. A laser Doppler
perfusion image analyzer (Moor Instruments) recorded changes in blood
flow postoperatively. Blood was collected by retro-orbital bleeding with the
use of heparinized capillaries, and white blood cells (WBCs) were counted.
Plasma samples were stored at ⫺30°C. Mice were killed 15 or 28 days after
resection of the femoral artery.
PBMC transplantation
Blood was collected from GFP mice with or without tPA treatment on days 0, 1,
and 2. PBMCs isolated from 400 ␮L of blood were injected intravenously
into HL ischemia–induced mice every day for a period of 3 days.
Isolation and transplantation of PB-derived CD11bⴙ and
VEGFR-1ⴙ cells
C57BL/6 male donor mice were treated with or without tPA from day 0 to
day 2. Blood was collected daily from each mouse. Cells were labeled with
anti-CD11b magnetic beads (Miltenyi Biotec) or biotinylated VEGFR-1
antibody (clone, MF-1; ImClone Systems), followed by avidin-Microbead
isolation by magnetic-activated cell sorting (Miltenyi Biotec). Isolated
CD11b⫹, CD11b⫺ VEGFR-1⫹, and VEGFR-1⫺ cells were greater than
90% pure by fluorescence-activated cell sorter (FACS) analysis. PB-derived
VEGFR-1⫹ and VEGFR-1⫺ cells (0.3 ⫻ 105 cells/injection/d) or CD11b⫹
and CD11b⫺ cells (1.7 ⫻ 105 cells/injection/d) from male donor mice were
transplanted daily intravenously into untreated HL ischemia–induced
C57BL/6 female mice for a period of 3 days. HL ischemia and cell
transplantation was started at the same time.
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OHKI et al
In vivo blocking experiments
HL ischemic mice, treated with or without rtPA (day 0-2; 10 mg/kg body
weight in 150 ␮L of 0.2% BSA) were coinjected intraperitoneally with
800 ␮g of anti–mouse VEGFR-1 (clone MF-1), anti–mouse VEGFR-2
(clone DC101), or control immunoglobulin G (IgG) on days 0, 2, and 4. HL
ischemic tPA-treated (day 0-2) and nontreated mice were coinjected with
500 ␮g of anti-CD11b antibody (clone 5C6) on days 0, 2, and 4 in the
presence or absence of VEGFR-1 and VEGFR-2 antibodies (see above).
tPA-treated (day 0-2) and nontreated mice were coinjected with 50 ␮g of
anti–mouse VEGF-A (R&D Systems) on day 0, or with 125 ␮g of c-Kit
(clone ACK; hybridoma kindly provided by S. Nishikawa, RIKEN, Kobe,
Japan) on days 0 and 2. Appropriate isotype antibody controls were
included.
ELISA
Plasma samples from tPA-treated and nontreated mice, with or without
induction of HL ischemia, were assayed for murine VEGF-A by enzymelinked immunoabsorbent assay (ELISA; R&D Systems).
In vitro culture supernatants of tPA-mobilized cells
PBMCs (2 ⫻ 105) isolated from tPA-treated and nontreated C57BL/6 mice
at different time points were cultured in 200 ␮L of serum-free medium for
24 hours. Supernatant analysis of VEGF-A was performed with ELISA.
Immunohistochemistry
Ischemic muscle tissue samples were cryopreserved with Tissue-Tek OCT
Compound (Sakura). Transverse-cut tissue samples of the whole leg were
prepared, serum-blocked, and stained with the first overnight at 4°C.
Serum-blocked sections were identified with the CD31 antibody (clone,
MEC13.3; PharMingen) followed by biotin-conjugated goat anti–rat IgG
(Vector) and Cy3-conjugated streptavidin (Alexa 594; Molecular Probes)
staining. Myelo-monocytic cells were identified with the VEGFR-1 (clone
MF-1; ImClone Systems) and CD11b antibody (clone M1/70; PharMingen)
followed by biotin-conjugated goat anti–rat IgG (Vector) and FITC- or
Cy3-conjugated streptavidin (Molecular Probes) staining. Nuclei were
counterstained with DAPI (Molecular Probes). In addition, tissues were
stained with antibodies against murine VEGF-A (clone A-20; Santa Cruz
Biotechnology), followed by goat anti–rabbit IgG conjugated with Alexa
594 (Molecular Probes). Immunoreactions were visualized with the use of a
Carl Zeiss fluorescence microscope. The capillary density (mean number of
capillaries per mm2) of CD31-stained slides was determined as described.22
Ischemic area (mean size per mm2) of H&E-stained slides was determined
with the use of computerized image analysis software (KS400; ⫻40
objective).
Statistical analyses
All data are presented as the mean plus or minus SEM. Intergroup
comparisons were performed by paired Student t test or by analysis of
variance. Probability values of P less than .05 were interpreted to denote
statistical significance.
Results
tPA mobilizes CD11bⴙ cells into the circulation, a process
dependent on plasmin and MMP-9–mediated release of Kit
ligand and VEGF-A
We observed that intraperitoneal or intravenous (data not shown)
injection of an engineered, serpin-resistant form of tPA or uPA (data
not shown) into C57BL/6 mice increased the number of WBCs
(Figure 1A), including neutrophils and monocytes (Figure 1B-C).
The absolute number of cells mobilized after tPA treatment was
lower than after treatment with the cell-mobilizing cytokine
BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
G-CSF. Further study will be needed to understand whether both
agents share a common downstream pathway during the mobilization process. No change after tPA administration was observed in
the number of CD45⫺ cells (data not shown). Within the CD45⫹
cell population, tPA augmented the number of CD45⫹CD11b⫹ cells
(Figure 1D), which were identified as CD45⫹CD11bhigh neutrophils, CD45⫹CD11bmed/F4/80med monocytes, and CD45⫹CD11blow/
F4/80⫺ cells (supplemental Figure 1A, available on the Blood Web
site; see the Supplemental Materials link at the top of the online
article). Treatment of mice with anti-CD11b prevented tPAinduced myeloid CD11b⫹ cell mobilization (data not shown),
supporting earlier studies that CD11b is required for monocyte/
neutrophil migration through the endothelium.23 CD45⫹CD11b⫹
cells isolated from tPA-treated but not BSA-PBS–treated mice
showed higher expression of the angiogenesis-associated genes
VEGF-A, CD184 (also known as CXCR-4), GM-CSF, CC chemokine receptor 2 (CCR2; a monocyte chemotactic protein-1 receptor), and neuropilin-124 by reverse transcription–PCR (supplemental Figure 1B) and CD184, and VEGFR-1 and c-Kit by FACS
analysis (supplemental Table 1). Tie-2 gene expression was only
detected on the CD45⫹CD11b⫺ cell fraction isolated from tPAtreated mice.25
The angiogenic factor VEGF can be liberated from ECM stores
after plasmin or MMP activation.16,26,27 We found that tPA administration augmented plasma VEGF-A levels in wild-type but not in
MMP⫺/⫺ and Plg⫺/⫺ mice or in mice treated with tranexamic acid,
a drug impairing tPA-mediated plasmin activation (Figure 1E-G).
Western blotting confirmed that plasma samples of tPA-treated
animals expressed 44-kDa biologically active VEGF-A (data not
shown). PBMCs isolated from tPA-treated animals at the indicated
time points released VEGF-A into culture supernatants (Figure
1H). Anti-VEGF treatment prevented tPA-mediated CD11b⫹ myeloid cell mobilization (Figure 1I), which is in accordance with
previous studies showing that VEGF-A can mobilize or promote
migration of hematopoietic VEGFR-1⫹, endothelial VEGFR2⫹28,29 and macrophages.30
We reported that tPA administration increases the number of
BM cells, a process partially dependent on KitL signaling.10 To
address the question of whether tPA-induced VEGF-A has an effect
on BM cell expansion, we coinjected tPA-treated C57BL/6 mice
with or without neutralizing antibodies against murine VEGF-A.
Blocking VEGF-A signaling with neutralizing antibodies against
murine VEGF-A did indeed prevent tPA-mediated BM cell expansion in vivo (data not shown).
CD11b/CD18 (␣M␤2, Mac-1) is a receptor for fibrin.31 tPA can
induce cell migration by binding to CD11b and degrading fibrin.5
We showed increased fibrinogen staining in BM cells in a model of
stress-hematopoiesis, indicating that fibrinogen might play a role in
hematopoietic cell retention and mobilization.10 Batroxobin, a drug
that reduces circulating fibrinogen, prevented the tPA-mediated
WBC (data not shown) and CD11b⫹ cell increase (Figure 1J).
However, further studies will be necessary to fully understand the
mechanism by which fibrinogen regulates cell mobilization. So
does tPA-mediated cell mobilization depend on plasmin activation?
Catalytically inactive tPA did not increase WBC (data not shown)
and CD11b⫹ cell counts (Figure 1K). Similarly, a drug impairing
tPA-mediated plasmin activation (Figure 1L) or genetic ablation of
Plg (Figure 1M) blocked the tPA-induced WBC (data not shown)
and CD11b⫹ cell increase. We noticed elevated baseline WBC
counts in Plg⫺/⫺ mice compared with Plg⫹/⫹ mice, but the
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BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
Figure 1. tPA mobilizes hematopoietic myelomonocytic cells into the circulation, a process dependent
on plasmin and MMP-9–mediated growth factor release. (A-D) C57BL/6 mice were injected intraperitoneally with recombinant tPA (n ⫽ 8 per group) or G-CSF
(n ⫽ 3) from day 0 until day 2. The total number of WBCs
(A), neutrophils (B), and monocytes (C) were counted.
(D) FACS analysis of CD45⫹CD11b⫹ cells in peripheral
blood of tPA-treated, G-CSF–treated, or BSA-PBS–
treated mice (n ⫽ 3 per group). (E-G) Plasma VEGF-A
levels were determined by ELISA in MMP-9⫺/⫺ (E), Plg⫺/⫺
(F) and wild-type control mice (n ⫽ 6) as well as in mice
treated with tranexamic acid, a drug that impairs plasmin
activation (n ⫽ 3), after tPA administration. Double asterisks indicate a significant difference between tPA-treated
MMP-9 or Plg wild-type mice and their matching knockout
controls. (H) PBMCs were isolated at the indicated time
points from tPA-treated and untreated C57BL/6 mice.
Cells were cultured under serum-free conditions for
24 hours. Supernatants were analyzed for VEGF-A by
ELISA (n ⫽ 6). (I) tPA-treated and untreated C57BL/6
mice were coinjected with antibodies against murine
VEGF-A or (J) cotreated with batroxobin intraperitoneally
(fibrinogen-reducing agent). (K) Mice were treated with
tPA, nonenzymatically active tPA, or vehicle control.
(L) tPA- or carrier-injected C57BL/6 mice were coinjected
with the plasmin inhibitor tranexamic acid. (I-L) The
frequency of circulating CD45⫹CD11b⫹ cells was determined on day 2 (n ⫽ 3 per group). (M) Plg⫹/⫹ and Plg⫺/⫺
mice received tPA intraperitoneally or BSA-PBS from day 0 to
day 2. WBC counts were assessed (n ⫽ 5 per group;
**P ⬍ .001; comparing Plg⫺/⫺ to Plg⫹/⫹ group). (N) C57BL/6
mice were injected with tPA and anti–c-Kit or control antibodies. The frequency of circulating CD45⫹CD11b⫹ cells was
measured on day 2 (n ⫽ 3 per group). (O) WBC counts were
determined in MMP-9⫹/⫹ and MMP-9⫺/⫺ mice after tPA
administration (n ⫽ 4). The asterisk indicates a significant
difference between tPA-treated MMP-9⫺/⫺ and MMP-9⫹/⫹
mice. (I-L,N) Asterisks denote a significant difference between the indicated groups. (A-D) Asterisks indicate a significant difference between the BSA-PBS and tPA or G-CSF
groups. Single asterisk, P ⬍ .05; double asterisk, P ⬍ .001.
Values represent the mean ⫾ SEM.
tPA-DRIVEN MYELOID CELL DRIVEN TISSUE REGENERATION
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BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
OHKI et al
difference was not statistically significant. These data indicate that
plasmin activation is required for cell mobilization.
KitL promotes hematopoietic cell survival, proliferation, and
mobilization.32 tPA administration can promote myeloid cell expansion by MMP-9–mediated release of KitL from stromal/niche
cells.10 Indeed, antibodies against c-Kit blocked tPA-mediated
CD11b⫹ cell mobilization (Figure 1N), and tPA-mediated cell
mobilization did not occur in MMP⫺/⫺ mice (Figure 1O). These
data indicate that c-Kit signaling is required for tPA-mediated cell
mobilization.
tPA administration accelerates ischemic revascularization
tPA is secreted after vascular injury. We exploited a HL ischemic
model to show that tPA was required for CD11b⫹ myeloid cell
mobilization, neoangiogenesis, and ischemic muscle tissue regeneration. Even though tPA⫺/⫺ mice showed normal CD11b⫹ cell
counts at baseline, kinetic studies showed enhanced numbers of
circulating CD11b⫹ cells in tPA⫹/⫹ but not in tPA⫺/⫺ mice during
the natural course of HL ischemic recovery (Figure 2A). Administration of tPA restored CD11b⫹ cell mobilization in HL ischemic
tPA⫺/⫺ mice similar to wild-type levels. Next, we studied whether
tPA accelerates HL ischemic recovery. Faster collateral vessel
growth (Figure 2B) and blood flow recovery (Figure 2C) were seen
in the ischemic limb of tPA-treated than in BSA-PBS–treated mice.
Ischemic tissue sections from tPA-treated wild-type mice showed
less fatty degeneration (Figure 2D), smaller areas of necrotic tissue
(Figure 2E), and higher capillary density (Figure 2F-G) than
controls. tPA augmented the number of ischemic muscle-residing
VEGF-A⫹ cells coexpressing VEGFR-1 (Figure 2H-J) in HL
ischemia–induced mice compared with controls. tPA improved
blood flow recovery and increased capillary density (data not
shown) in tPA⫺/⫺ mice (Figure 2K) but was ineffective in Plg⫺/⫺
(Figure 2L) or MMP-9⫺/⫺ mice (data not shown). In summary
these data showed that endogenous tPA plays a role during the
natural course of ischemic recovery, which coincided with impaired CD11b⫹ cell mobilization. In addition, exogenous tPA
administration accelerates ischemic tissue regeneration and
neoangiogenesis.
Ex vivo tPA-treated CD11bⴙ and VEGFR-1ⴙ cells improve
neoangiogenesis
To determine the proangiogenic potential of tPA-mobilized cells,
we isolated PBMCs from GFP donor mice receiving tPA or vehicle
and transplanted them daily into HL ischemic recipients. Mice that
received a transplant with tPA-mobilized PBMCs showed faster
blood flow recovery (Figure 3A), smaller areas of fatty degeneration (Figure 3B), and increased capillary density (Figure 3C)
compared with mice injected with BSA-PBS–stimulated PBMCs.
Nearly all donor-derived GFP⫹ cells coexpressed CD11b (Figure
3D-F) and were detectable in ischemic muscle tissues of mice
injected with tPA-mobilized PBMCs but not in tissues of mice
injected with control cells.
To examine whether CD11b⫹ cells, known to enhance both
normal and malignant neoangiogenesis,33 were the effector cells for
tPA-driven neoangiogenesis, we transplanted equal numbers of
tPA-mobilized CD11b⫹ cells into HL recipient mice. tPAmobilized but not BSA-PBS–mobilized CD11b⫹ cells accelerated
ischemic reperfusion (Figure 3G) and increased capillary density in
ischemic tissues of HL ischemic recipients (Figure 3H). Antibodies
against CD11b prevented tPA-mediated HL ischemic tissue recovery and blocked tPA-driven vessel formation (Figure 3I-J). tPA not
only augmented the absolute number of circulating CD11b⫹ cells
(supplemental Table 2), but tPA-stimulated cells on a cellular basis
also improved tissue regeneration better than BSA-PBS–stimulated
CD11b⫹ cells. Because approximately 50% of CD11b⫹ cells
coexpressed VEGFR-1 (supplemental Table 1), and the frequency
of CXCR4⫹VEGFR-1⫹ cells/hemangiocytes was augmented after
tPA therapy compared with controls (1.08% ⫾ 0% versus
0.195% ⫾ 0.005%, respectively; n ⫽ 3 per group; P ⬍ .05), we
examined the effect of tPA on VEGFR-1–driven neoangiogenesis.
Transplanted tPA-stimulated VEGFR-1⫹ cells improved tissue
ischemic regeneration (Figure 3K). Ischemic tissue revascularization was achieved with a 5-fold lower numbers of tPA-stimulated
VEGFR-1⫹ cells than tPA-stimulated CD11b⫹ cells. These data
suggest that tPA-mediated neoangiogenesis is driven by a
CD11b⫹VEGFR-1⫹ cell population.
Inhibition of VEGF-A signaling prevents tPA-mediated cell
mobilization and tissue neoangiogenesis
Plasmin can release VEGF-A from ECM stores.26 tPA administration in HL ischemic animals resulted in VEGF-A plasma elevation
(Figure 4A). We performed reciprocal transplants of wild-type
hematopoietic cells into lethally irradiated Plg⫺/⫺ mice or vice
versa (Plg⫹/⫹ BM cell transplantation into Plg⫺/⫺ recipients or
Plg⫺/⫺ BM cell transplantation into Plg⫹/⫹ recipients) followed by
induction of HL ischemia. We show that tPA treatment increased
plasma VEGF-A levels in HL Plg wild-type mice with or without
transplantation of Plg wild-type BM cells (Figure 4B). Plasma
VEGF-A levels were impaired in HL ischemic Plg-deficient
recipient mice. These data implied that plasmin activation within
the nonhematopoietic environment was required for tPA-mediated
systemic VEGF-A elevation (see also Figure 1E-G). Within
ischemic tissues, we found VEGF-A staining mainly in CD11b⫹
cell by immunohistochemistry (Figure 4C). VEGF-A administration
partially rescued impaired angiogenesis observed in Plg⫺/⫺ HL ischemic
mice, as reported,34 but did not in MMP-9⫺/⫺mice (Figure 4D-H).
But is VEGF-A signaling required for tPA-mediated tissue
neoangiogenesis? Blockade of VEGF signaling with antibodies
against murine VEGF-A (Figure 5A-B) or VEGF (Figure 5C-F)
receptors inhibited tPA-mediated ischemic tissue recovery and
neoangiogenesis as well as myeloid cell mobilization by VEGFR-1
(Figure 5G-I) in an HL ischemic model. Similarly, tPA did not
improve neoangiogenesis in myeloid cell–depleted mice (Figure
5J). Coinjection of anti–VEGFR-1 antibodies into myeloid cell–
depleted mice further reduced neoangiogenesis both in BSA-PBS–
and tPA-treated mice. These data indicate that CD11b⫹ hematopoietic cells are the main “angiogenic effector cells” for tPA-mediated
cell-driven neovascularization. However, it is possible that some
non-CD11b/VEGFR-1⫹ cells (eg, endothelial cells) might contribute to the tPA-induced angiogenic response.
Discussion
There is convincing evidence that a small number of angiogenic
cells are present in the blood of most mammals, including humans.
In the present study, we demonstrate that plasmin activation
through administration of serpin-resistant mutant tPA alone was
sufficient to mobilize BM-derived hematopoietic, angiopotent cells
into the circulation. Our findings support a mechanism whereby
tPA-generated plasmin mobilizes myeloid cells through the release
of VEGF-A and KitL, processes that are dependent on MMP-9
activation and, possibly, on that of other MMPs.
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tPA-DRIVEN MYELOID CELL DRIVEN TISSUE REGENERATION
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Figure 2. tPA administration accelerates revascularization in a HL ischemia model. (A) tPA or vehicle was
injected intraperitoneally daily from day 0 to day 2 into
tPA⫹/⫹ and tPA⫺/⫺ mice after HL ischemia induction. The
frequency of circulating CD45⫹CD11b⫹ cells was measured (n ⫽ 3 per group). The single dagger (†) symbol
denotes a significant difference (P ⬍ .05) between day
0 and day 2 in HL tPA⫹/⫹ mice. (B) C57BL/6 mice were
treated with tPA or vehicle. Macroscopic pictures of the
lower limb region were taken on day 15 after HL induction. Images were acquired with 0.5⫻ and 2.5⫻/0.1
numeric aperture (NA) lenses (corresponding to 5⫻ and
25⫻ magnification), an Olympus SZX7 camera, and
Adobe Photoshop CS3. (C) The ischemic/normal blood
flow ratio, determined by analysis of the whole crosssection of the limb by laser Doppler perfusion imaging
analysis, is shown as a function of time. (D-G) Evaluation
of necrotic areas (D-E, after 15 days; bar ⫽ 200 ␮m) and
capillary density in sections (F-G, after 28 days;
bar ⫽ 50 ␮m) of HL-induced mice treated with or without
tPA (n ⫽ 6 per group). Panels D and F were acquired with
a 20⫻/0.50 NA and a Carl Zeiss KS400 camera, and
analyzed using Axio Vision (Carl Zeiss). (H-I) Quantification of VEGF-A⫹ cells (arrows) in ischemic tissues (n ⫽ 6
per group, after 15 days; bar ⫽ 20 ␮m). Images were
acquired with a 40⫻/0.75 NA and a Carl Zeiss KS400
camera, and analyzed using Axio Vision (Carl Zeiss).
(J) Immunohistochemical staining of VEGFR-1 and
VEGF-A of ischemic muscle tissues derived from vehicleor tPA-treated HL ischemic mice and nonischemic mice.
Arrows indicate VEGFR-1⫹VEGF-A⫹ cells (bar ⫽ 20 ␮m).
Images were acquired with a 40⫻/0.75 NA and a Carl
Zeiss KS400 camera, and analyzed using Axio Vision
(Carl Zeiss). (K-L) tPA or vehicle was injected intraperitoneally daily into tPA (K) or Plg (L) wild-type and deficient
mice after induction of HL ischemia. Functional perfusion
measurements of the collateral region were performed
(n ⫽ 3 per group). A significant difference between tPA⫹/⫹
and tPA⫺/⫺ mice treated with BSA-PBS is shown. If not
otherwise mentioned, asterisks denote significant differences between BSA-PBS– and tPA-treated groups.
*P ⬍ .05; **P ⬍ .001; values represent the mean ⫾ SEM.
The number of circulating cells increasingly used for autologous and allogeneic transplants and tissue regeneration can be
significantly improved with cytokines such as G-CSF.22 Stressinduced hematopoietic cell egress from the BM is a multistep
process, involving activities of cytokines and chemokines, as well
as proteolytic enzymes. Proliferation and/or differentiation in the
BM usually precedes enhanced hematopoietic cell egress.35 We
recently demonstrated that tPA alone had no effect on myeloid
precursor cell expansion in vitro, but increased BM cell numbers in
vivo/in the presence of stromal cells a process partially dependent
on c-Kit/KitL signaling.10 Here, we report that the tPA-mediated
increase in BM cells (myeloid cells) also requires VEGF-A
signaling, because neutralizing antibodies against VEGF-A prevented tPA-mediated BM cell (including CD11b⫹) cell expansion
(data not shown). VEGF-A is a principal regulator of blood vessel
formation and hematopoiesis and is capable of mobilizing hematopoietic progenitors and differentiated myeloid cells.28,36 In addition, the c-Kit/KitL signaling pathway is also implicated in cell
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BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
Figure 3. Transplantation of tPA-treated CD11bⴙ and
VEGFR-1ⴙ myeloid cells improve neoangiogenesis.
(A-F) tPA- or BSA-PBS–mobilized PBMCs from GFP
mice or carrier (no cells) were injected intravenously into
HL ischemia–induced recipients. (A) Blood flow was
determined. (B-F) Ischemic tissues of lower limbs isolated 15 days after injection and HL ischemia induction
were stained with (B) H&E and (C) antibodies against
CD31 for capillary density evaluation (n ⫽ 3 per group).
Images were acquired with 20⫻/0.50 NA and a Carl Zeiss
KS400 camera, and analyzed using Axio Vision (Carl
Zeiss). (D-E) Within ischemic muscle tissues GFP⫹ donor
cells (arrows) were quantified. Inserts depict a magnified
view. Images were acquired with a 20⫻/0.50 NA and a
63⫻/1.3 oil objective using a Carl Zeiss KS400 camera.
(E) Quantification of the number of GFP⫹ cells and GFP⫹
cells coexpressing CD11b in ischemic tissues (n ⫽ 8 per
group). (F) CD11b (Mac-1) staining of lower limb ischemic
tissue from mice receiving carrier injections, from mice
that received a transplant with PBMCs, or from BSA-PBS–
treated or tPA-treated donor GFP mice. Arrows indicate
transplanted GFP⫹CD11b⫹ cells in ischemic tissues
(bar ⫽ 20 ␮m). Images were acquired with 40⫻/0.75 NA
and a Carl Zeiss KS400 camera, and analyzed using Axio
Vision (Carl Zeiss). (G-H) PB-derived CD11b⫹ or CD11b⫺
cells isolated from tPA- or BSA-PBS–treated donors were
transplanted into HL ischemia–induced recipients for
3 days (n ⫽ 4 per group). (G) Blood flow was determined.
(H) Capillary density was evaluated. (I-J) HL ischemic
C57BL/6 mice were treated with tPA and coinjected with
anti-CD11b or control monoclonal antibodies. (I) Blood
flow was analyzed by laser Doppler analysis. (J) Capillary
density was evaluated. (K) PB-derived VEGFR-1⫹ or
VEGFR-1⫺ cells, isolated from tPA- or BSA-PBS–treated
donors, were injected into HL ischemia–induced recipients, and blood flow was examined (n ⫽ 4 per group).
Single asterisk (P ⬍ .05) and double asterisk (P ⬍ .001)
indicate a significant difference between the groups.
Values represent mean ⫾ SEM.
migration/mobilization.37 Here, we demonstrate that tPA administration increases circulating proangiogenic CD45⫹CD11b⫹ cells
coexpressing both VEGFR-1 and c-Kit, a process which could be
blocked with the use of antibodies against VEGF-A and c-Kit.
Interestingly, intraventricular leukocytosis has been observed in
rats with intraventricular hemorrhage/hematoma that were treated
with high doses of tPA.38 Streptokinase administration in patients
with acute myocardial infarction caused a marked increase in
circulating WBCs.39
MMP-9 has been shown to be required for macrophage
migration and is induced in premetastatic lung endothelial and BM
cells, including macrophages after VEGF-A treatment involving
VEGFR-1.29,40 Here, we report that tPA-mediated CD11b⫹ cell
mobilization requires the presence of MMP-9. MMP-9, which is
produced in the BM, eg, after treatment with, eg, G-CSF,37 is able
to degrade occludin, one of the components that comprise the
endothelial tight junctions.41 Monocytes can induce tPA release in
brain endothelial cells, which subsequently activates extracellular
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tPA-DRIVEN MYELOID CELL DRIVEN TISSUE REGENERATION
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Figure 4. Administration of recombinant VEGF-A
partially rescues the angiogenic defect observed in
Plg- and MMP-9–deficient mice. (A-B) Plasma VEGF-A
levels were analyzed in tPA-treated and untreated HLinduced C57BL/6 (A) and chimeric (B) mice (n ⫽ 3 per
group). Chimerism was achieved as follows: Lethally irradiated Plg⫹/⫹ or Plg⫺/⫺ mice received a transplant with Plg⫹/⫹
or Plg⫺/⫺ donor BM cells. Six weeks later, HL ischemia–
induced chimeric mice were treated intraperitoneally
with/without tPA. Asterisks and single dagger indicate a
significant difference between tPA-treated and BSA-PBS–
treated groups. (C) Immunohistochemical staining for
CD11b and VEGF-A of ischemic muscle tissue derived
from tPA-treated and vehicle-treated HL ischemic mice.
Arrows indicate CD11b⫹ VEGF-A⫹ cells (bar ⫽ 25 ␮m).
Images were acquired with 40⫻/0.75 NA and a Carl Zeiss
KS400 camera, and analyzed using Axio Vision (Carl
Zeiss). (D-H) HL-induced wild-type control (D-E,H), Plg⫺/⫺
(F,H), and MMP-9⫺/⫺ (G,H) mice were injected intraperitoneally daily with tPA, mVEGF-A, or BSA-PBS from day 0
until day 2. (D) Functional perfusion measurements of the
collateral region with the use of a laser scan at the
indicated time points in wild-type animals. Representative
laser Doppler perfusion images are shown of the regions
analyzed. Blood perfusion (red to yellow) was augmented
in tPA-treated mice compared with vehicle-injected
controls (green to blue ⫽ reduced perfusion signal).
(E-G) Blood flow recovery was determined by laser
Doppler perfusion imaging in the ischemic limbs (n ⫽ 3;
*P ⬍ .05, **P ⬍ .001). (H) Capillary density was evaluated on CD31-stained sections 15 days after ligation
(n ⫽ 3; **P ⬍ .001). Asterisks indicate a significant difference between the indicated groups. Error bars represent
SEM.
signal–regulated kinase 1/2. Both tPA and extracellular signal–
regulated kinase 1/2 have been shown to control the breakdown of
the tight junction protein occludin.17 Determination of the extent to
which the integral plasma membrane protein occludin mediates
some of the tPA-induced myeloid cell mobilization by modulating
the BM endothelial barrier awaits further studies.
The mechanism by which tPA stimulates angiogenesis is
unclear. The angiogenic effect of tPA was seen as early as 5 to
7 days after surgery. These data suggest that the kinetics of
angiogenic cell mobilization by tPA may mirror that observed for
hematopoietic cell mobilization peaking on day 2. Here, we
demonstrate that the angiogenic effect of tPA can be adoptively
transferred by blood mononuclear cells. Of interest, tPA administration increased not only the absolute number of these angiopotent
cells, but also more importantly tPA-stimulated cells showed
qualitatively improved angiogenic performance/potential compared with BSA-PBS–treated controls in an HL ischemic murine
model. This is in contrast to the reported proangiogenic effects of
other mobilizing agents such as G-CSF, which cause an increase in
the basal numbers of circulating mononuclear cells rather than
inducing the mobilization of a unique angiogenic cell population.42
Within the mobilized cell population, we identified a tPAresponsive, hematopoietic CD45⫹CD11b⫹VEGFR-1⫹, angiopotent and readily transplantable cell population, which may serve as
a basis for tPA therapy in patients with ischemia. tPA-mobilized
CD45⫹CD11b⫹ cells also showed higher gene expression of
neuropilin-1, but not Tie-2, which has been shown to indicate a
more angiogenic/arteriogenic cell population within the mononuclear cell fraction.24,25 Monocytes are key contributors to angiogenesis at sites of ischemic injury because they can secrete
proangiogenic cytokines, including VEGF-A, and basic fibroblast
growth factor.43
VEGF-A is a critical survival factor for vascular endothelium.44 The role of plasmin in the ischemic, proteolytic microenvironment can function as a determinant controlling VEGF-A–
mediated activities. VEGF189 or VEGF206, proteins bound to
extracellular heparin-containing proteoglycans, can be released
by plasmin, and the released 34-kDa dimeric species can
function as endothelial cell mitogens.26 However, recent data
have shown that the angiogenic potential of matrix-associated
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OHKI et al
BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
Figure 5. Inhibition of VEGF signaling prevented
tPA-mediated ischemic tissue recovery. (A-B) C57BL/6
mice were treated with tPA and coinjected with anti–
VEGF-A or control monoclonal antibodies. (A) Ischemic
recovery was quantified by laser Doppler (n ⫽ 3 per
group). Asterisks indicate a significant difference between tPA-treated mice coinjected with IgG or anti–
VEGF-A antibody. (B) Capillary density was quantified
15 days after ligation (n ⫽ 3 per group). HL ischemic mice
treated with or without tPA were coinjected intraperitoneally with neutralizing doses of VEGFR-1 (C), VEGFR-2
(D), a combination of VEGFR-1 and VEGFR-2 (E), or IgG
antibodies at 2-day intervals, starting from day 0, and
functional perfusion measurements of the collateral region were performed in each group with the use of a laser
scan (n ⫽ 3; *P ⬍ .05,**P ⬍ .001). (F) Quantification of
the capillary density in tissue sections taken 15 days after
artery ligation by CD31 staining of sections from HLinduced mice treated with or without tPA (n ⫽ 3; *P ⬍ .05;
**P ⬍ .001 versus vehicle). (G-I) Blood cell counts were
determined at the indicated time points. (G) WBC,
(H) neutrophil, and (I) monocyte counts (*P ⬍ .05 comparison of the tPA plus IgG-treated HL group with the
BSA-PBS plus IgG-treated group; †P ⬍ .05 comparison
of the tPA plus IgG-treated HL group with the tPA plus
anti–VEGF-A group; #P ⬍ .05 comparison of the tPA plus
IgG-treated HL group with the tPA plus anti–VEGFR-1
group). Error bars represent SEM. (J) HL-induced
C57BL/6 mice treated with tPA or BSA-PBS and antibodies against CD11b were cotreated with antibodies against
VEGFR-1 or VEGFR-2. Capillary density was evaluated
(n ⫽ 3 per group). If not otherwise indicated, a single
asterisk (P ⬍ .05) and double asterisk (P ⬍ .001) indicate a significant difference between groups. Values
represent the mean ⫾ SEM.
VEGF-A isoforms is superior to soluble isoforms.45 We showed
that tPA administration results in systemic VEGF-A plasma
elevation, which required plasmin activation, because genetic
ablation of Plg and pharmacologic inhibition of plasmin activation reduced systemic VEGF-A elevation. We showed that the
extent of revascularization after tPA administration correlates
with absolute plasma VEGF-A levels and increased numbers of
tissue-residing CD11b⫹ cells coexpressing VEGF-A. Neutralizing antibodies against VEGF receptors or VEGF-A in vivo
abolished tPA-mediated tissue regeneration.
Proteolysis of fibrin matrices by endothelial cells plays
essential roles in the migratory and morphogenic differentiation
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BLOOD, 27 MAY 2010 䡠 VOLUME 115, NUMBER 21
tPA-DRIVEN MYELOID CELL DRIVEN TISSUE REGENERATION
processes underlying angiogenesis. Because fibrin is part of the
provisional matrix needed to support vascular cell responses for
repair, the binding of growth factors and cytokines, such as
VEGF-A, will localize them to the site of injury so that they can
enhance the cellular responses needed for angiogenesis.46
Thus, activation of the fibrinolytic cascade promotes both
mobilization of hematopoietic cells and their incorporation into
the ischemic tissue. This hematopoietic cell recruitment is a
critical step in regulating the supply of local and systemic
angiogenic factors, which finally accelerates ischemic revascularization. Because VEGF-A also increases the expression and
activity of PAs, especially tPA in endothelial cells,47 based on
our data, we propose that tPA production in endothelial cells is
required for the recruitment of hematopoietic cells to neoangiogenic sites, including peripheral ischemic sites. These data are
consistent with our findings that disruption of the tPA gene
reduced the number of CD11b⫹ cells during the natural course
of HL ischemic recovery and with data of other groups showing
impaired macrophage migration in tPA⫺/⫺ mice during peritonitis and sciatic nerve injury.48,49 Macrophage migration during
inflammation has been shown to depend on CD11b/Mac-1
recognition of a binary complex consisting of fibrin within the
provisional matrix and tPA.5 Cao et al5 elegantly demonstrated
that subsequent neutralization of tPA by its inhibitor plasminogen activator inhibitor-1 enhanced binding of the integrinprotease-inhibitor complex to the endocytic receptor lipoprotein
receptor–related protein, triggering a switch from cell adhesion
to cell detachment.
Our combined data show that a serpin-resistant form of tPA
induces leukocyte mobilization and promotes neoangiogenesis.
Our findings have implications for regenerative medicine: Administration of tPA alone or in combination with other growth factors
might be a novel strategy to increase the efficiency of hematopoietic, angiopotent cell harvests for cell-based therapy of ischemic
diseases. These data set forth the novel concept that plasmin
activation, apart from controlling coagulation, controls myeloid
cell–driven neoangiogenesis during tissue regeneration.
4311
Acknowledgments
We thank Dr A. Furuhata and S. Nakamura from Juntendo
University and the FACS core facility for their help, and ImClone
(NY) for providing antibodies against murine VEGF receptors.
Human recombinant tPA was a generous gift from Eisai Corporation (Japan). Pauline O’Grady, a medical writer, provided editorial
assistance to the authors during the preparation of this manuscript.
This work was supported by research grants from the Japan Society
for the Promotion of Science and Grants-in-Aid for Scientific Research
from Ministry of Education, Culture, Sports, Science and Technology
(K.H. and B.H.); in part by research grants from the Ministry of Health,
Labour and Welfare (K.H.), Mitsubishi Pharma Research Foundation
(K.H.), Kyowa Hakko Kirin Co Ltd. (K.H,), Japan Leukaemia Research
Fund (K.H,), Novartis Foundation for the Promotion of Science (B.H.),
the Sagawa Foundation for Promotion of Cancer Research (B.H.), Kato
Memorial Bioscience Foundation (B.H.), Astellas Foundation for Research on Metabolic disease (B.H.); by grants from the Lundbeck
Foundation (L.R.L.) and the National Cancer Institute (Z.W.). This work
was supported by a Program for Improvement of the Research
Environment for Young Researcher (B.H.) funded by the Special
Coordination Funds for Promoting Science and Technology of the
Ministry of Education, Culture, Sports, Science and Technology, Japan.
Authorship
Contribution: Y.O., M.O., L.R.L, H.N., K.H., Z.W., and B.H. wrote
the manuscript; L.R.L, Z.Z., H.O., Z.W., K.O., H.Y, and A.N. supplied
the reagents; and K.Y., Y.O., M.O., C.N., M.I., Y.T., K.H., H.A., H.K.
and B.H. designed and performed experiments, and analyzed data.
Conflict-of-interest disclosure: The authors declare no competing financial interests.
Correspondence: Koichi Hattori, Center for Stem Cell Biology
and Regenerative Medicine, Institute of Medical Science, University of Tokyo, 4-6-1, Shirokanedai, Minato-ku, Tokyo 108-8639,
Japan; e-mail: [email protected].
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From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
2010 115: 4302-4312
doi:10.1182/blood-2009-08-236851 originally published
online January 28, 2010
Tissue type plasminogen activator regulates myeloid-cell dependent
neoangiogenesis during tissue regeneration
Makiko Ohki, Yuichi Ohki, Makoto Ishihara, Chiemi Nishida, Yoshihiko Tashiro, Haruyo Akiyama,
Hiromitsu Komiyama, Leif R. Lund, Atsumi Nitta, Kiyofumi Yamada, Zhenping Zhu, Hideoki Ogawa,
Hideo Yagita, Ko Okumura, Hiromitsu Nakauchi, Zena Werb, Beate Heissig and Koichi Hattori
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