The WNT antagonist Dickkopf2 promotes angiogenesis in rodent and human endothelial cells.
ABSTRACT Neovessel formation is a complex process governed by the orchestrated action of multiple factors that regulate EC specification and dynamics within a growing vascular tree. These factors have been widely exploited to develop therapies for angiogenesis-related diseases such as diabetic retinopathy and tumor growth and metastasis. WNT signaling has been implicated in the regulation and development of the vascular system, but the detailed mechanism of this process remains unclear. Here, we report that Dickkopf1 (DKK1) and Dickkopf2 (DKK2), originally known as WNT antagonists, play opposite functional roles in regulating angiogenesis. DKK2 induced during EC morphogenesis promoted angiogenesis in cultured human endothelial cells and in in vivo assays using mice. Its structural homolog, DKK1, suppressed angiogenesis and was repressed upon induction of morphogenesis. Importantly, local injection of DKK2 protein significantly improved tissue repair, with enhanced neovascularization in animal models of both hind limb ischemia and myocardial infarction. We further showed that DKK2 stimulated filopodial dynamics and angiogenic sprouting of ECs via a signaling cascade involving LRP6-mediated APC/Asef2/Cdc42 activation. Thus, our findings demonstrate the distinct functions of DKK1 and DKK2 in controlling angiogenesis and suggest that DKK2 may be a viable therapeutic target in the treatment of ischemic vascular diseases.
- SourceAvailable from: Stefan Liebner[Show abstract] [Hide abstract]
ABSTRACT: The blood-brain barrier (BBB) is essential for maintaining homeostasis within the central nervous system (CNS) and is a prerequisite for proper neuronal function. The BBB is localized to microvascular endothelial cells that strictly control the passage of metabolites into and out of the CNS. Complex and continuous tight junctions and lack of fenestrae combined with low pinocytotic activity make the BBB endothelium a tight barrier for water soluble moleucles. In combination with its expression of specific enzymes and transport molecules, the BBB endothelium is unique and distinguishable from all other endothelial cells in the body. During embryonic development, the CNS is vascularized by angiogenic sprouting from vascular networks originating outside of the CNS in a precise spatio-temporal manner. The particular barrier characteristics of BBB endothelial cells are induced during CNS angiogenesis by cross-talk with cellular and acellular elements within the developing CNS. In this review, we summarize the currently known cellular and molecular mechanisms mediating brain angiogenesis and introduce more recently discovered CNS-specific pathways (Wnt/β-catenin, Norrin/Frizzled4 and hedgehog) and molecules (GPR124) that are crucial in BBB differentiation and maturation. Finally, based on observations that BBB dysfunction is associated with many human diseases such as multiple sclerosis, stroke and brain tumors, we discuss recent insights into the molecular mechanisms involved in maintaining barrier characteristics in the mature BBB endothelium.Cell and Tissue Research 03/2014; · 3.33 Impact Factor
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ABSTRACT: Objective Ovarian hyperstimulation syndrome is associated with increased angiogenesis and vascular leakage. Immature myeloid cells (IMCs) and dendritic cells have been shown to be actively involved in angiogenesis in several disease modells in mice and humans. Nevertheless, little is known about the role of these cells in the ovary. As such, this study sought to determine whether alterations in these ovarian myeloid cell populations are associated with gonadotropin stimulation in a mouse modell. Study design Four-week-old pre-pubertal C57Bl/6 female mice were allocated into three groups: high-dose stimulation (n = 4; pregnant mare serum gonadotropins (PMSG) 20 U for 2 days), low-dose stimulation (n = 5; PMSG 5 U for 1 day) and sham-treated controls (n = 4). Human chorionic gonadotropin 5 U was injected on Day 3, and the mice were killed on Day 5. Ovaries were analysed by flow cytometry, confocal microscopy and quantitative polymerase chain reaction. Results Gonadotropin stimulation increased the proportion of CD11b+Gr1+ IMCs among the ovarian myeloid cells: 22.6 ± 8.1% (high dose), 7.2 ± 1.6% (low dose) and 4.1 ± 0.3% (control) (p = 0.02). Conversely, gonadotropin stimulation decreased the proportion of ovarian CD11c+MHCII+ dendritic cells: 15.1 ± 1.9% (high dose), 20.7 ± 4.8% (low dose) and 27.3 ± 8.2% (control) (p = 0.02). IMCs, unlike dendritic cells, were localized adjacent to PECAM1+ endothelial cells. Finally, gonadotropin stimulation was associated with increased expression of S100A8, S100A9, Vcan and Dmbt1, and decreased expression of MMP12. Conclusions Gonadotropin stimulation is associated with proangiogenic myeloid cell alterations, reflected by a dose-dependent increase in ovarian IMCs and a parallel decrease in dendritic cells. Recruited IMCs localize strategically at sites of angiogenesis. These changes are associated with differential expression of key proangiogenic genes.Fertility and Sterility 08/2014; · 4.30 Impact Factor
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ABSTRACT: The adult mammalian heart predominantly comprises myocytes, fibroblasts, endothelial cells, smooth muscle cells and epicardial cells arranged in a precise three dimensional framework. Following cardiac injury the spatial arrangement of cells is disrupted as different populations of cells are recruited to the heart in a temporally regulated manner. The alteration of the cellular composition of the heart after cardiac injury thus enables different phenotypes of cells to interact with each other in a spatio-temporal dependent manner. It can be argued that the integrated study of such cellular interactions rather than the examination of single populations of cells can provide more insight into the biology of cardiac repair especially at an organ wide level. Many signaling systems undoubtedly mediate such cross talk between cells after cardiac injury. The Wnt/β-catenin system plays an important role during cardiac development and disease and here we describe how cell populations in the heart after cardiac injury mediate their interactions via Wnt/β-catenin pathway, determine how such interactions can affect a cardiac repair response and finally suggest an integrated approach to study cardiac cellular interactions.Cardiovascular Research 03/2014; · 5.81 Impact Factor
1882? The?Journal?of?Clinical?Investigation? ? ? http://www.jci.org? ? ? Volume 121? ? ? Number 5? ? ? May 2011
The WNT antagonist Dickkopf2
promotes angiogenesis in rodent
and human endothelial cells
Jeong-Ki Min,1,2 Hongryeol Park,1 Hyun-Jung Choi,1 Yonghak Kim,1 Bo-Jeong Pyun,1,3
Vijayendra Agrawal,1 Byeong-Wook Song,4 Jongwook Jeon,5 Yong-Sun Maeng,1
Seung-Sik Rho,1 Sungbo Shim,6 Jin-Ho Chai,1 Bon-Kyoung Koo,7 Hyo Jeong Hong,2
Chae-Ok Yun,8 Chulhee Choi,5 Young-Myoung Kim,9 Ki-Chul Hwang,4 and Young-Guen Kwon1
1Department of Biochemistry, College of Life Science and Biotechnology, Yonsei University, Seoul, Republic of Korea.
2Therapeutic Antibody Research Center, Korea Research Institute of Bioscience and Biotechnology, Daejeon, Republic of Korea.
3College of Pharmacy and Division of Life Science and Pharmaceuticals, Ewha Womans University, Seoul, Republic of Korea.
4Cardiovascular Research Institute, Yonsei University College of Medicine, Seoul, Republic of Korea. 5Cell Signaling and Bioimaging Laboratory,
Department of Bio and Brain Engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon, Republic of Korea.
6Department of Biological Science, Sookmyung Women’s University, Yongsan-Ku, Seoul, Republic of Korea. 7Department of Biological Sciences,
Seoul National University, Silim-dong, Gwanak-gu, Seoul, Republic of Korea. 8Brain Korea 21 Project for Medical Science,
Institute for Cancer Research, Yonsei Cancer Center, Yonsei University College of Medicine, Seoul, Republic of Korea.
9Vascular System Research Center, Kangwon National University, Kangwon-Do, Republic of Korea.
Angiogenesis, the formation of new blood vessels from a preexist-
ing vascular bed, is essential for embryonic development as well as
many pathophysiological processes in adult life, including wound
healing, diabetic retinopathy, tumor growth, and metastasis (1–3).
During angiogenesis, specialized ECs located at the vascular front
undergo dynamic and coordinated phenotypic alterations in mor-
phology, cell-to-cell (and/or cell-to-matrix) communication, and
proliferation (4–6). These cellular changes are accompanied by
the concomitant dynamic regulation of specific gene expression
within a developing vessel, which is indispensable for establishing
a functional vascular network (4, 5).
Numerous studies have identified many genes with localized
expression at the vascular front and delineated their specific func-
tions in regulating EC sprouting and remodeling during develop-
ment of the vascular tree (4, 5). VEGF, the most well-characterized
angiogenic molecule, has been shown to guide endothelial tip cell
filopodia via VEGFR2 at the sprouting front (2, 7). Receptors of
neuronal axon guidance cues such as Unc5b, EphB4, Plexin D1,
and Robo4 are also prominently expressed at the vascular front
and play significant roles in vessel path finding and sprouting
(8–10). Recently, Notch signaling has been shown to be a key
player in vascular morphogenesis, particularly in confining tip
cell and stalk cell characters, in addition to arterial-venous deter-
mination (5). Notch1 binding by Delta-like 4 (Dll4) suppresses
tip cell formation (11–13), and blockade of Dll4-Notch1 signal-
ing in mice inhibits tumor growth by increasing the formation of
nonproductive vessels with excessive sprouting and many small
vessel branches (14, 15). Benedito et al. further showed that Jag-
ged1 (Jag1) antagonizes Dll4-Notch signaling in ECs expressing
Fringe family glycosyltransferases and promotes angiogenesis
in mice (16). The identification and characterization of factors
regulating EC type specification and dynamics should be valu-
able for further understanding of the molecular mechanisms of
neovascularization and for establishing new strategies for treat-
ing angiogenesis-dependent diseases.
WNT pathways have been widely implicated in regulating the
development and maintenance of the vascular system (17). A large
number of studies have demonstrated diverse roles of WNT signal-
Authorship?note: Jeong-Ki Min and Hongryeol Park contributed equally to this
Conflict?of?interest: The authors have declared that no conflict of interest exists.
Citation?for?this?article: J Clin Invest. 2011;121(5):1882–1893. doi:10.1172/JCI42556.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
ing in regulating EC functionalities such as proliferation, survival,
differentiation, cell junctions, and polarity (18). Indeed, genetic
studies have demonstrated that WNT 7b induces hyaloid vessel
regression (19) and Norrin-Frizzled4 signaling plays a key role in
the vascular development of the eye and ear (20, 21). WNT signals
are also considered to be orchestrated with other pathways, includ-
ing Notch and TGF-β, to regulate vascular morphogenesis (18).
However, the function of WNTs and the underlying regulatory
pathways in the vasculature remain unclear due to the complexity
in spatial expression and interaction of large number of WNTs and
their regulatory proteins.
In the present study, we observed that Dickkopf1 (DKK1) and
DKK2, originally known as WNT antagonists that bind to LDL
receptor–related protein (LRP) 5/6 and inhibit β-catenin signaling
pathways (22–24), are differentially expressed in ECs during mor-
phogenetic differentiation on Matrigel. By employing biochemi-
cal and Tg analysis, we demonstrate that DKK1 and DKK2 play
opposite functions in regulating angiogenesis. Importantly, DKK2
was found to significantly improve tissue repair with functional
neovascularization in ischemic disease models. Furthermore, this
study provides what we believe to be novel mechanistic insights
regarding DKK2 in stimulating angiogenic sprouting.
Reciprocal expression of DKK1 and DKK2 during endothelial morphogenesis.
To identify factors potentially acting at the sprouting front, we
employed Affymetrix oligonucleotide arrays (GEO GSE27160)
to isolate genes that reversibly change their expression during in
vitro proliferation and morphogenesis of ECs. Interestingly, we
detected reciprocal change in the expression of Dickkopf fam-
ily members DKK1 and DKK2. DKK1 was rapidly downregulated
upon induction of morphogenesis on 2D Matrigel, whereas DKK2
was upregulated during this process (Figure 1A). These changes
in expression were reversible (Supplemental Figure 1; supple-
mental material available online with this article; doi:10.1172/
JCI42556DS1) and transcriptionally (Figure 1B) controlled during
morphogenesis on Matrigel and proliferation on gelatin.
We further investigated the regulation of DKK1 and DKK2 gene
expression. None of the angiogenic factors tested nor hypoxia had
an effect on DKK1 and DKK2 expression (Supplemental Figure 2,
A and B). However, DKK2 was sufficiently induced by EC recogni-
tion of laminin, a major component of Matrigel, and did not require
morphological changes in ECs (Supplemental Figure 2, C–E).
In contrast, downregulation of DKK1 was correlated with the induc-
tion of morphological changes in ECs (Supplemental Figure 2,
DKK1 and DKK2 reciprocally expressed during endothelial morphogenesis distinctively regulate angiogenesis in vitro. (A) mRNA and protein
levels of DKK1 and DKK2 at the same time points (0.5, 8, and 12 hours) during morphogenesis in Matrigel were measured by RT-PCR (left)
and Western blotting (right). (B) Human DKK1 or DKK2 promoter activities were measured by luciferase reporter assay. HUVECs cultured on
gelatin-coated plates were transfected and transferred to gelatin-coated plates (gelatin) or Matrigel-coated plates (morphogenesis) 6 hours later.
Luciferase reporter activity was measured 16 hours later. Data represent mean ± SD. (C) HUVECs stably expressing eGFP plus control (CTL)
shRNA, eGFP plus DKK1 shRNA, or DKK1 plus control shRNA were established with lentivirus. Stable transfectants were plated on Matrigel-
coated plates at a density of 1.5 × 105 cells/well and incubated for 18 hours. Capillary-like networks, which completely differentiated into tube-like
structure, were quantified with Image-Pro Plus software. (D) Proliferative indices of HUVECs transfected with eGFP or DKK1 were assessed
by [3H]-thymidine incorporation assay. (E) HUVECs stably expressing eGFP, DKK2, control shRNA, or DKK2 shRNA. Morphogenesis of the
transfectants on Matrigel was analyzed as described in C. (F) Proliferative indices of HUVECs transfected with eGFP or DKK2 were accessed
as described in D. Data represent mean ± SD. **P < 0.01; ***P < 0.001.
1884?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
F and G) and was not significantly affected by the recognition of
specific ECM components (Supplemental Figure 2C).
Distinct roles of DKK1 and DKK2 during in vitro angiogenesis. We next
determined the roles of DKK1 and DKK2 in angiogenesis in vitro.
Sustained expression of DKK1 during morphogenesis or treatment
with recombinant DKK1 protein sup-
pressed the formation of EC tube-like
structures on 2D Matrigel (Figure 1C
and Supplemental Figures 3 and 5).
In contrast, overexpression of DKK2
or treatment with recombinant DKK2
protein enhanced the integrity of EC
tube-like structures, while knockdown
of DKK2 during morphogenesis inhib-
ited it (Figure 1E and Supplemental
Figures 4 and 5). In addition, DKK2
treatment of HUVEC-coated beads
caused more sprouts of HUVECs com-
pared with controls on fibrin bead assay
(Supplemental Figure 6). Interestingly,
EC proliferation on gelatin was also
significantly inhibited by DKK1 overex-
pression and increased by DKK1 shRNA
transfection (Figure 1D and Supple-
mental Figure 7). Overexpression of
DKK2 had no significant effect on EC
proliferation on gelatin (Figure 1F).
These results suggest that DKK1 and
DKK2 perform opposite functions dur-
ing angiogenesis in vitro.
DKK2 is a proangiogenic factor in vivo.
To evaluate the effects of DKK1 and
DKK2 on angiogenesis in vivo, we
employed the Matrigel implant assay
in mice. DKK2- or VEGF-containing
Matrigel plugs were red in color, imply-
ing neovascularization (Supplemental
Figure 8). The color of Matrigel plugs
cotreated with VEGF and DKK1 was
similar to that of control-treated plugs
(Supplemental Figure 8). Quantifica-
tion of hemoglobin and CD31 staining
of Matrigel plugs consistently indicat-
ed that DKK2 significantly increases
angiogenesis compared with controls,
while DKK1 suppresses VEGF-induced
angiogenesis (Figure 2, A and B).
We further examined the structural
characteristics of DKK2-induced blood
vessels in a mouse cornea pocket assay.
As previously demonstrated (25), the
growth of VEGF-induced blood ves-
sels was relatively fast, but their struc-
ture appeared to be disorganized and
leaky (Figure 2C). In contrast, DKK2
induced well-organized vascular net-
works with distinct vascular tree-like
structures (Figure 2C). In addition,
Evans blue administration demon-
strated that DKK2-induced vessels are
less leaky than VEGF-induced vessels (Figure 2D). Compared
with VEGF-stimulated capillaries, DKK2-induced vessels con-
sistently showed more coverage of ECs by pericytes and SMCs,
which play an important role in vessel maturity and stability
(Figure 2, E and F, and Supplemental Figure 9).
DKK2 protein induces angiogenesis in vivo. (A and B) Matrigel plugs treated with VEGF (200 ng)
and DKK2 (1 μg) were excised from mice 5 days after injection (n = 5 per group). Scale bars: 100 μm.
CD31 staining (A) and quantification of hemoglobin concentrations in a Matrigel plug (B). White
arrows indicate CD31-stained vessels. (C–F) Corneal angiogenic responses induced by VEGF-
containing (200 ng) or DKK2-containing (1 μg) micropellets. Scale bars: 500 μm. Asterisks indi-
cate micropellet inserted (C). 5 days after pellet implantation, Evans blue was injected i.v. After
30 minutes, each mouse eye was photographed. Scale bars: 200 μm (D). CD31 and NG-2 staining
of cryosections of the eye (E). Scale bars: 100 μm. Green, CD31 positive; red, NG-2 positive; blue,
DAPI. Pericyte coverage calculated as a ratio of the NG-2 to CD31 staining area (F). Data represent
mean ± SD. ***P < 0.001.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
DKK2 Tg mice exhibit increased vessel formation in the retina. (A–F) Isolectin B4 staining of whole-mounted P4 retinas of WT or DKK2 Tg
mice (A and D). White broken lines indicate the range of vessel extension (A). Scale bars: 500 μm. White dots indicate filopodia extended
from tip cells (D). Scale bars: 100 μm. Quantification of vessel densities (B), sprouting length (C), tip cell number (E), and filopodia (F). (G–I)
Isolectin B4 staining of whole-mounted P12 retinas (G). Scale bars: 100 μm. Quantification of vessel density in the ganglion (first) (H) and the
outer nuclear (third) cell layer (I). (J–M) Aortic segments were harvested from WT and DKK2 Tg mice (n = 7 per group). (J) Endothelial sprouts
forming branching cords from the margins of aortic segments were photographed with a phase microscope. Scale bars: 200 μm. (L) Dynamic
movement of endothelial sprouts from the margins of aortic segments was captured as real-time video (see Supplemental Video 1). Scale bars:
40 μm. Arrows indicate filopodia. Sprouting scores (K) and quantification of tip cell numbers (M). Sprouting scores were scored from 0 (least
positive) to 5 (most positive). Data represent mean ± SD. *P < 0.05; ***P < 0.001. Box indicates 25%~75% value and whisker indicates media
value in box plot (B, C, E, F, H, and I).
1886?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
Enhanced angiogenic sprouting and filopodial dynamics in DKK2 Tg
mice. To further examine the angiogenic function of DKK2 in vivo,
we generated Tg mice that express mDKK2 under the control of
the EC-specific Tie2 promoter/enhancer (Supplemental Figure 10).
In the 3 Tg mouse lines generated, we could not detect significant
developmental abnormalities at the embryonic stage or in adults.
However, DKK2 Tg embryos generally showed slightly increased
capillary density at E9.5 and beyond compared with WT litter-
mates (data not shown). Since the growth rate and patterning of
retinal blood vessels during early development are well character-
ized, we analyzed the retinal phenotype in DKK2 Tg mice. At P4,
DKK2 Tg mice displayed increased vascular density and sprout-
ing length in retinal vasculature compared with WT littermates
(Figure 3, A–C). The number of tip cells and filopodia were also
consistently increased in DKK2 Tg mice (Figure 3, D–F). Ki-67
staining also showed increased endothelial cell proliferation in
DKK2 Tg retinas compared with WT littermates (Supplemental
Figure 11). The discrepancy between previous in vitro results and
current in vivo results may be due to the difference between in
vitro culture and in vivo growth conditions. For pericyte recruit-
ment, DKK2 Tg mice did not appear to differ from WT littermates
(Supplemental Figure 11). In P12 retinas, the vascular density,
internal filopodia, and branching of the ganglion layer (first layer)
as well as the vascular plexus in the deeper layer (third layer) were
markedly increased in DKK2 Tg mice (Figure 3, G–I, and Supple-
mental Figure 12). Further, the increase in retinal vascular den-
sity was maintained up to 10 weeks after birth (Supplemental
Figure 13). In an ex vivo aortic ring assay, the average length and
branch number of endothelial sprouts were consistently increased
in DKK2 Tg aortas compared with those from WT littermates
(Figure 3, J and K, and Supplemental Figure 14). In time-lapse
video microscopy recorded over 9 hours, DKK2 Tg aortas showed
highly dynamic filopodial protrusion of tip cells (Figure 3, L and M,
and Supplemental Video 1).
DKK2 promotes new vessel formation in both hind limb ischemia and
myocardial infarction animal models. We next examined the effect of
local injection of DKK2 protein on therapeutic neovasculariza-
tion in a murine model of hind limb ischemia. Hind limb isch-
emia was induced by ligation and excision of the right femoral
artery and vein, resulting in a severe vascular perfusion defect
(26). Intramuscular injection of DKK2 resulted in a significant
reduction in necrosis compared with ischemic mice treated
with PBS (Figure 4, A and B). Indocyanine green (ICG) imag-
ing revealed that blood perfusion was significantly improved
at 3 days after surgery in mice injected with DKK2 compared
with PBS-injected mice (Figure 4, A and B), consistent with the
hypothesis that the reduced necrosis was due to improved neo-
vascularization in the ischemic limb.
We further assessed the therapeutic effect of DKK2 in a rat
model of myocardial infarction (MI). One week after MI induction
and DKK2 protein injection, the size of the LV infarct was evalu-
ated in the DKK2 and control (sham) groups. Injection of DKK2
protein significantly decreased the infarct size and the degree of
fibrosis in the infarct zone compared with sham (Figure 4, C–F).
The mean microvessel count per field in the infarcted heart was
also higher (Figure 4, G and H), and the incidence of TUNEL-posi-
tive myocardial cells was significantly reduced in the DKK2 group
compared with sham (Figure 4, I and J).
Changes in cardiac function in DKK2 protein–treated MI ani-
mals were measured with transthoracic echocardiography. Echo-
cardiography data (Figure 4K, Supplemental Figure 15, and
Supplemental Table 1) showed that DKK2 increased both LV
fractional shortening (FS) and LV ejection fraction (EF) compared
with sham. Injection of DKK2 or VEGF showed further improve-
ment in systolic performance and cardiac dimensions compared
with sham animals. However, cardiac dimensions including LV end
diastolic diameter (LVEDD) and LV end systolic diameter (LVESD)
were smaller in the DKK2-treated group than in the VEGF-treated
group (Supplemental Figure 15 and Supplemental Table 1). The
DKK2 group had a higher increase in systolic performance (16.27%
increment in percentage of FS and 20.5% increment in percentage
of EF) than the VEGF group (Figure 4K, Supplemental Figure 15,
and Supplemental Table 1).
DKK2-induced filopodial extrusion of ECs requires Cdc42 activation. We
next scrutinized the mechanism of action of DKK2 in the vascula-
ture. Previous studies showed that tip cell filopodial extension in
the developing retina depends on astrocyte-derived VEGF-A acting
directly on tip cells (7, 8). However, glial fibrillary acidic protein
(GFAP) and VEGF-A mRNA levels, as well as the relative levels of
VEGF-A splice isoforms, remained unchanged in DKK2 Tg mice
compared with WT mice (Supplemental Figure 16), suggesting a
direct action for DKK2 in ECs.
The tips of the sprouts are composed of highly migratory cells
with numerous filopodia- and lamellipodia-like processes, which
are conferred by the action of small GTPases, such as Cdc42 (27).
Therefore, we hypothesized that DKK2 may stimulate tip cell
filopodial extension through regulating the activation of Cdc42.
In HUVECs cultured on gelatin, overexpression of DKK2 but
not DKK1 induced Cdc42 activation (Figure 5A). Consistently,
both overexpression of DKK2 and treatment with DKK2 protein
induced Cdc42 activation in retinal endothelial cells (RECs) (Sup-
DKK2 promotes angiogenesis and improves tissue recovery in animal
models of hind limb ischemia and MI. (A and B) Intramuscular injec-
tion of DKK2 increased blood perfusion and reduced the probability
of necrosis in the ischemic hind limb in mice. Tissue perfusion rate
(%/min) was defined as the fraction of blood exchanged per minute in
the vascular volume by time-series analysis of indocyanine green dye.
Blood perfusion rate of the hind limb was measured at postoperative
day 0 (POD 0). Correlation between regional perfusion rates of the
ischemic hind limbs at POD 0 and limb necrosis levels at POD 3 was
determined. The x axis shows the regional perfusion rate of the isch-
emic hind limbs (poor perfusion rate: 0%~30%/min; moderate perfusion
rate: 30%~100%/min; good perfusion rate: 100 < %/min). Normal hind
limbs typically demonstrate a perfusion rate above 400%/min. (C–J)
Increase of myocardial repair after DKK2 injection. Myocardial injection
of DKK2 decreased the LV infarct size as assessed by TTC staining
at 1 week after MI (C and D). Representative images taken from a
Masson’s trichrome–stained section (muscle is stained red, collagen
is stained blue) (E and F). Microvessel staining with CD31 (G) and the
TUNEL assay (I) on cardiac muscle tissues. Scale bars: 50 μm. Quan-
titative analysis of microvessel density and the TUNEL assay (H and
J). Area with yellow broken lines (C) indicates the infarct region. Red
arrows (G and I) indicate microvessels and apoptotic cells, respec-
tively. (K) Effective improvement of cardiac function by DKK2 injection.
Cardiac functions were measured with 2D conventional parameters:
FS and LVEF at 3 weeks after injection of DKK2 into MI rats. FS (%) =
([LVEDD – LVESD]/LVEDD) × 100 (%). LVEDV = 7.0 × LVEDD3/(2.4
+ LVEDD), LVESV = 7.0 × LVESD3/(2.4 + LVESD), and LVEF (%) =
(LVEDV – LVESV)/LVEDV × 100. *P < 0.05; **P < 0.01; ***P < 0.001.
S; Sham, V; VEGF, D2; DKK2. Data represent mean ± SD.
1888?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
plemental Figure 17). During HUVEC morphogenesis, Cdc42 was
spontaneously activated in HUVECs plated on Matrigel and its
activation was slightly potentiated in DKK2-expressing HUVECs,
but significantly reduced in DKK1-expressing cells (Figure 5B
and Supplemental Figure 18). In addition, HUVEC and REC filo-
podial protrusion was significantly increased by treatment with
DKK2 protein at 2 hours compared with controls, and a domi-
nant negative form of Cdc42 blocked DKK2-induced filopodial
extensions of HUVECs (Figure 5, C and D, and Supplemental
Figures 17 and 19). These results suggest that DKK2 may pro-
mote angiogenesis, at least in part, by endowing the dynamics
of tip cell structures, which require Cdc42 activation and filopo-
dial protrusion. To identify potential supporting mechanisms of
DKK2-mediated Cdc42 activation in ECs, we determined whether
the WNT/Frizzled complex or LRP5/6 is involved in the signaling
events induced by DKK2. Treatment with soluble Frizzled-relat-
ed protein (sFRP), a decoy antagonist of WNT, did not inhibit
DKK2-mediated Cdc42 activation (Figure 5E).
Interestingly, knockdown experiments of LRP5 and LRP6
revealed that LRP6, but not LRP5, was critically involved in DKK2-
mediated Cdc42 activation, although both LRPs are expressed in
ECs (Figure 6A and Supplemental Figure 20, A–C). Consistently,
siRNA specific to LRP6 completely blocked DKK2-induced EC
morphogenesis, whereas siRNA against LRP5 had no significant
effect (Figure 6, B and C, and Supplemental Figure 15, D and E).
LRP6 expression was also specifically detected in retinal ECs (Sup-
plemental Figure 21). These results suggest that DKK2 activates
the Cdc42 signaling pathway via LRP6 and independent of WNT/
It has recently been reported that adenomatous polyposis coli
(APC) stimulates the guanine nucleotide exchange factor (GEF) activ-
ity of APC-stimulated exchange factor 2 (Asef2), a guanine-nucleo-
tide exchange factor specific for Cdc42 (28). Since APC is one of the
components working downstream of LRP6, we hypothesized that
DKK2 stimulation of LRP6 might induce APC/Asef2 complex for-
mation, resulting in Cdc42 activation. Indeed, APC and Asef2 inter-
DKK2 increases filopo-
dial protrusions in a Cdc42-
dependent manner. (A)
HUVECs stably expressing
eGFP, DKK1, or DKK2 were
analyzed for Cdc42 activities.
(B) Stable transfectants were
cultured on Matrigel-coated
plate (morphogenesis) for
2 hours and Cdc42 activi-
ties were measured. (C and
D) HUVECs were transiently
transfected with expression
plasmids encoding eGFP
control or eGFP dominant
negative mutant of Cdc42
(Cdc42 N17). After 24 hours,
these cells were plated on
Matrigel-coated plates and
incubated with PBS or DKK2
(1.5 μg/ml) for 2 hours. Then
microphotographs were taken
(C) and filopodia number was
quantified (D). Arrowheads
indicate filopodial extension.
Data represent mean ± SD.
**P < 0.01. Scale bars: 20 μm.
(E) HUVECs stably express-
ing eGFP or DKK2 were incu-
bated with sFRP (200 ng/ml)
for 24 hours and Cdc42 activ-
ity was measured.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
action was prominently increased in ECs expressing DKK2 and was
blocked by LRP6 siRNA (Figure 6, D and E). Moreover, knockdown
of APC or Asef2 inhibited the Cdc42 activation and EC morphogen-
esis induced by DKK2 (Figure 6, F–H, and Supplemental Figure 22).
Taken together, these results indicate that the angiogenic action of
DKK2 may be mediated through its cognate receptor LRP6 in ECs
and that at least the APC/Asef2 complex lies downstream of LRP6,
leading to Cdc42 activation and stimulation of tip cell dynamics.
In this study, we propose different biological roles of DKK1 and
DKK2 in regulating angiogenesis. Expression of DKK1 and DKK2
was reciprocally and specifically regulated depending on the sta-
tus of ECs, such as proliferation and morphogenetic differentia-
tion. DKK2 promoted angiogenesis, whereas DKK1 suppressed it.
DKK1 and DKK2 have similar primary structures and were origi-
nally identified as soluble WNT antagonists (29). These proteins
regulate various biological functions through inhibition of the
WNT signaling pathway (22–24). Interestingly, DKK2 is also sug-
gested to work as an activator depending on cell context (29, 30).
However, the biological function of DKK2 as an LRP5 or LRP6
agonist in vivo and the underlying signaling mechanism remain
to be determined. Our finding that DKK2 promotes angiogenesis
provides a new insight into the agonist function of DKK2 in vivo.
DKK2-induced Cdc42 activation requires LRP6-mediated APC/Asef2 signaling. (A–C) HUVECs stably expressing eGFP or DKK2 were tran-
siently transfected with control, LRP5-specific, or LRP6-specific siRNA. After 60 hours, Cdc42 activity was measured (A). Cells were plated on
Matrigel-coated plates at a density of 1.5 × 105 cells/well and incubated for 18 hours. Microphotographs were taken and capillary-like networks
were quantified with Image-Pro Plus software (B and C). (D) Coimmunoprecipitation of APC with Asef2. Lysates were prepared from cells
overexpressing eGFP or DKK2, immunoprecipitated with anti-APC antibody, and probed with anti-Asef2 or anti-APC antibodies. (E) eGFP- or
DKK2-expressing cells were transiently transfected with control or LRP6-specific siRNA. After 60 hours, cell lysates were subjected to coim-
munoprecipitation. (F–H) eGFP- or DKK2-expressing cells were transiently transfected with control, APC-specific, or Asef2-specific siRNA. After
60 hours, Cdc42 activity was measured (F). The cells were plated on Matrigel-coated plates at a density of 1.5 × 105 cells/well and incubated for
18 hours. Capillary-like networks were quantified with Image-Pro Plus software (G and H). Data represent mean ± SD. ***P < 0.001.
1890?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
Our data suggest that the angiogenic activity of DKK2 is cor-
related with activation of EC dynamics. The vascular-specific
DKK2 Tg mice exhibited significantly increased vessel branch-
ing, EC density, and proliferation, with significantly enhanced
numbers of tip cells and filopodia at the vascular front of the
retina (Figure 3). Interestingly, these retinal vascular phenotypes
of DKK2 Tg mice were comparable to those of Jag1 Tg mice as
described previously (16). Since our study is based on the gain
of function, it is limited to arguing whether DKK2 is involved
in tip cell and stalk cell specification as Jag1 is. However, it is
most likely that the angiogenic sprouting and vascular pattern-
ing governed by DKK2 might be correlated with its stimulatory
function in tip cell dynamics, as clearly indicated in aortic ring
imaging (Figure 3, J–M, and Supplemental Video 1). Our data
showed that DKK2 is rapidly induced by EC recognition of lami-
nins, major components of the basement membrane. Notably,
it has been shown that laminins were produced by stalk cells
and tip cells during sprouting angiogenesis (31) and that lam-
inin-1 induces expression of Jag1 in ECs (32). We also observed
that both Jag1 and DKK2 are induced in a similar way during
morphogenesis of HUVECs on Matrigel (data not shown). Thus,
the expression pattern and Tg phenotypes point to a potential
relationship between Jag1 and DKK2 in sprouting angiogenesis.
Recent in silico analysis suggested that DKK2 is a Notch signal-
ing target in intestinal stem cells (33). However, Notch-response
elements were detected in the promoter region of DKK2 of
human, chimpanzee, and rat, but not in cow and mouse (33).
Thus, it remains unclear whether angiogenic action of DKK2 is
functionally correlated with Notch signaling.
This study further provides insight into the mechanism by
which DKK2 promotes sprouting angiogenesis. Filopodia extend-
ing from tip cells lead the angiogenic sprout by sensing guidance
cues and migrating along a specific path in response (7). Previous
studies have shown that Rho GTPases represent an important class
of molecules involved in vascular morphogenesis (34, 35). DKK2,
but not DKK1, consistently induced Cdc42 activation in ECs.
Although cultured ECs expressed both LRP5 and LRP6, which are
potential signaling receptors for DKK2, depletion of LRP6, but not
LRP5, resulted in a significant reduction in DKK2-induced Cdc42
activation and EC morphogenesis. LRP5 and LRP6 share 73% iden-
tity in protein sequences but the ligand-binding repeats show only
50% similarity (36), which suggests that they may not bind ligands
with similar affinity. Indeed, it has been shown that LRP6 works
differently from LRP5 in the WNT signaling pathway. Overexpres-
sion of LRP6, but not LRP5, induces axis duplication in Xenopus
(37), and only LRP6 cooperates with several WNT-Frizzled fusion
proteins to activate WNT-responsive promoters (38). Similarly,
our data suggest that the response of LRP5 and LRP6 to DKK2 in
ECs is distinct and LRP6 may act as a cognate DKK2 receptor that
mediates Cdc42 activation and angiogenic sprouting.
We further asked how the association of LRP6 with DKK2
activates Cdc42. APC is known to be involved in WNT signaling
downstream of LRP6 through its binding to β-catenin (39). Recent
studies have uncovered an additional function for APC, in which it
activates Cdc42 via direct binding to Asef2 (28, 40). Interestingly,
APC/Asef2 complex formation was enhanced by DKK2, and the
reduction of APC or Asef2 expression inhibited DKK2-induced
Cdc42 activation and EC morphogenesis, suggesting functional
significance of the APC/Asef2 system in linking DKK2/LRP6 to
Cdc42 activation in ECs. Some WNT members are known to acti-
vate Cdc42 via noncanonical pathways, promoting cell polarity
(41). However, treatment with sFRP, a decoy antagonist of WNT,
did not inhibit DKK2-induced Cdc42 activation (Figure 5E) and
DKK2 treatment did not increase β-catenin–mediated transcrip-
tion levels (Supplemental Figure 23), suggesting that DKK2 may
activate Cdc42 independent of the WNT pathway.
Norrin/LRP5/Fzd4 signaling is well known in regulating reti-
nal angiogenesis. EC-specific knockout mice of Norrin or Fzd4
showed reduced retina vessel growth and endothelial-mural cell
interaction (42). However, blocking of Norrin/LRP5/Fzd4 signal-
ing in vitro by Fzd4 CRD IgG did not interrupt Cdc42 activation
by DKK2. In addition, Cdc42 activation was not affected by Norrin
treatment (Supplemental Figure 24). In summary, CDC42 activa-
tion by the DKK2/LRP6 pathway does not seem to be affected by
Our data also demonstrate an inhibitory function of DKK1 in
angiogenesis with 2 possible mechanisms. First, unlike the WNT-
independent mechanism of DKK2 in vascular morphogenesis,
DKK1 appears to directly affect WNT signaling in ECs. Previous
studies have demonstrated roles for the WNT signaling pathway
in angiogenesis and vascular stability (18, 43). Signals transmit-
ted from the Frizzled family of receptors upon binding of its
ligands, WNT and Norrin, influence angiogenic processes such as
EC proliferation and migration (17). Moreover, mice deficient in
Lef1 and Ctnnb1, both of which control transcription of target
genes of the canonical WNT signaling pathway, displayed signifi-
cant reduction in vascular density in retinas (44). Our data show
that DKK1 inhibits WNT-responsive promoter activity and pro-
liferation of ECs (Figure 1D and Supplemental Figure 25), and
endothelium-specific overexpression of DKK1 in mice results in
delayed peripheral vascularization and reduced vascular density in
retinas (Supplemental Figure 26). Alternatively, DKK1 may coun-
teract DKK2 action through competitive binding to LRP6. This
mechanism is supported by our data showing that upregulation
of DKK1 antagonizes DKK2-induced angiogenic sprouting and
Cdc42 activation (Supplemental Figure 26).
Although knockout of DKK1 and DKK2 showed unique pheno-
types in head formation and osteoblast differentiation (45, 46),
respectively, no obvious vascular phenotype in mutant mice has
been reported so far. This may be due to the compensation for
DKK1 and DKK2 deficiency by other members of DKK family
(47) or other regulatory factors during normal vascular devel-
opment. And also, considering the complexity and diversity of
blood vessels in different organs and pathological states, care-
ful reexamination of vascular phenotypes in DKK-deficient mice
may be valuable for further understanding the precise roles of
DKKs in the vasculature.
The feature of DKK2-induced vessels, which were structurally
organized and well covered by pericytes and SMCs, was differ-
ent from those induced by VEGF as shown by the corneal assay.
DKK2 also provided profound therapeutic effects in 2 ischemic
disease models mentioned above. Although the discovery of sev-
eral angiogenic factors has led to the development of therapeutic
strategies for ischemic diseases, the success of therapeutic angio-
genesis in clinical trials has been limited due to insufficient effi-
cacy and pronounced adverse effects (48–51). Given that DKK2
is a relatively small and soluble ligand, our finding that DKK2
alone can induce new and functional vessel formation in vivo may
provide a novel strategy for developing therapeutic strategies to
treat ischemic vascular diseases.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
Cell culture. HUVECs were isolated from human umbilical cord veins by col-
lagenase treatment, as described previously (52), and used from passages
2–4. HUVECs were cultured in EC basal medium (EBM) or EC growth
medium (EGM, Clonetics) supplemented with 3% FBS.
In vitro morphogenesis on 2D Matrigel and oligonucleotide microarrays. In vitro
morphogenesis was assayed as previously described (53). Briefly, 800 μl
of growth factor–reduced Matrigel (BD Biosciences) was transferred to a
60-mm tissue culture well and polymerized for 30 minutes at 37°C.
HUVECs were plated onto the layer of Matrigel at 1 × 106 cells/plate and
cultured for 0.5, 8, or 18 hours. At each time point, total RNA was isolated
and hybridized to the HG-U133A 2.0 microarray (54675 human genes;
Affymetrix). The standard protocol used for sample preparation and
microarray processing is available from Affymetrix. Full data sets are avail-
able online (Gene expression analysis during morphogenesis of Human
umbilical vein endothelial cells; GEO GSE27160).
EC proliferation assay. HUVECs were seeded at 2 × 104 cells/well in gel-
atin-coated 24-well plates. After 48 or 72 hours, cells were washed twice
with M199 and incubated for 6 hours in M199 containing 0.5 μCi/ml of
[3H]thymidine (Amersham). High–molecular weight DNAs were precipi-
tated using 10% trichloroacetic acid at 4°C for 30 minutes. After 2 washes
with ice-cold H2O, 3H-radioactivity was solubilized in 0.2 N NaOH/0.1%
SDS and determined by liquid scintillation counter.
In vivo Matrigel plug assay. The Matrigel plug assay was performed as previ-
ously described. Briefly, 7-week-old C57BL/6 mice (Orient Co.) were injected
subcutaneously with 0.6 ml of Matrigel containing the indicated amount of
DKK2, DKK1, VEGF+DKK1, and 10 units of heparin. The injected Matrigel
rapidly formed a single, solid gel plug. After 5 days, the skin of the mouse
was easily pulled back to expose the Matrigel plug, which remained intact.
Hemoglobin was measured by the Drabkin method with Drabkin Reagent
Kit 525 (Sigma-Aldrich) to quantify blood vessel formation. The concentra-
tion of hemoglobin was calculated from a known amount of hemoglobin
assayed in parallel. To identify infiltrating ECs, immunohistochemistry was
performed with anti–CD-31 antibody (BD Biosciences).
Mouse corneal pocket assay. Eight-week-old C57BL/6 mice were anesthe-
tized with zoletil 50 (15 mg/kg) and lumpun (5 mg/kg). After 10 minutes,
alcaine was dropped into the eye. With a von Graefe cataract knife, a micro-
pocket was made in the cornea. Micropellets (0.35 × 0.35 mm) of sucrose
octasulfate-aluminum complex (s0652; Sigma-Aldrich) coated with poly-
2-hydroxyethyl methacrylate (p3932; Sigma-Aldrich) containing 200 ng of
VEGF (K0921632; KOMABIOTECH) and 1 μg of DKK2 were implanted
into the corneal micropocket. The pellet was positioned 0.6–0.8 mm from
the corneal limbus. Tardomyocel-L ointment was applied to the eye after
implantation. Eyes were examined with a microscope 7 days after implan-
tation. Cryosections were stained with anti-CD31 (BD Pharmingen) and
anti-NG2 (Millipore) antibodies.
Aortic ring assay. Aortas were harvested from 6- to 8-week-old C57BL/6
WT and DKK2 Tg mice. Plates (48-well) were coated with 100 μl Matrigel,
and after it had gelled, the rings were placed in the wells and sealed in place
with an overlay of 40 μl Matrigel. EGM was added to the wells in a final vol-
ume of 200 μl of human endothelial serum-free medium (Invitrogen). On
day 5, cells were fixed and stained with isolectin B4. The assays were scored,
double-blinded, from 0 (least positive) to 5 (most positive). Each data point
was assayed 6 times. Endothelial sprouts were time-lapse imaged on an
Olympus IX81-ZDC inverted fluorescence microscope (Olympus).
Analysis of mouse retinal vasculature. The vascular pattern of dissected
mouse retina was analyzed using a modification of a method previously
reported (7). Mice were sacrificed by ketamine injection and eyes were
enucleated in PBS. The eyes were fixed in 4% PFA-PBS (pH 7.4) for 1 hour
at 4°C. Retinas were dissected as described (54), postfixed in 70% ethanol
overnight at 4°C, washed with PBS, and permeabilized with PBS contain-
ing 1% Triton X-100 for 1 hour. Retinas were then incubated in blocking
solution for 4 hours at 37°C, followed by overnight incubation in Alexa
Fluor 488–conjugated Isolectin GS-IB4 solution (Molecular Probes) at 4°C.
After 5 washes in PBS containing 1% Triton X-100, the retinas were flat
mounted on slides using fluorescent mounting medium. Flat-mounted
retinas were analyzed by fluorescence microscopy using an Olympus IX81-
ZDC inverted fluorescence microscope and confocal fluorescent micro-
scope (LSM 510 META; Carl Zeiss).
Cryosection immunofluorescence staining. Tissue was fixed in 4% PFA-PBS
(pH 7.4) overnight at 4°C and rinsed with PBS at room temperature. Tissue
was incubated in 15% sucrose solution overnight at 4°C and then trans-
ferred to 30% sucrose at 4°C until the tissue sank. All fixation, rinsing,
and incubation were done gently. Tissue was infiltrated in OCT embed-
ding medium (Tissue Tek) for 0.5 hours at room temperature before freez-
ing. Tissue was transferred to an embedding mold, which was filled with
OCT. The mold was cooled with liquid nitrogen. After the material had
frozen, tissue was wrapped in aluminum foil and stored at –70°C. Sections
(8- to 50-μm thick) were cut at –20°C, and slides were stored at –70°C
until needed. Sections were prefixed in acetone for 0.5 hours at –70°C and
then dried briefly until the acetone was removed. OCT was removed with
water. Sections were incubated in blocking solution for 4 hours at 37°C or
overnight at 4°C, followed by overnight incubation in primary antibody at
4°C. After 5 washes in 0.3% Triton X-100 in PBS for 15 minutes each, the
sections were incubated in secondary antibody overnight at 4°C. Before
washing, the sections were treated with DAPI (1 μg/ml) and washed 5 more
times with 0.3% Triton X-100 in PBS for 15 minutes each. All antibodies
were dissolved in antibody diluent (Dako). Sections were analyzed by fluo-
rescence microscopy using an Olympus IX81-ZDC inverted fluorescence
microscope or a confocal microscope (LSM 510 META; Carl Zeiss).
Murine hind limb ischemia model. BalB/cAnNCriBgi-nu nude male mice were
obtained from Charles River Japan. All mice were 7–8 weeks (15–20 g) of age
at the time of study. Hind limb ischemia was induced by ligation and exci-
sion of the right femoral artery and vein under ketamine-xylazine anesthe-
sia. For angiogenic potential studies, mice were divided into 3 groups after
induction of ischemia. Mice received intramuscular injection of PBS (20 μl
with 0.9% NaCl) alone as a control, saline solution containing DKK2 (20 μl
of 600 ng/μl), or saline solution containing VEGF (20 μl of 150 ng/μl).
NIR fluorescence imaging. We used customized optical systems for NIR
fluorescence imaging, as previously described by Kang et al. (55). Briefly,
this system employs a CCD digital camera (PIXIS 1024; Princeton Instru-
ments) with a custom-made 830-nm band-pass filter (Asahi Spectra USA)
and 760-nm light-emitting diode arrays (SMC760; Marubeni America). For
time-series ICG imaging, mice under ketamine-xylazine anesthesia were
injected with an i.v. bolus of ICG (0.1 ml of 400 mM/l; Sigma-Aldrich) into
the tail vein. ICG fluorescence images were obtained in a dark room for
10 minutes at 1-second intervals immediately after injection. After serial
imaging, customized computer programs were used to obtain perfusion
maps and necrosis probabilities.
MI model and experimental procedure. MI was induced in 8-week-old
Sprague-Dawley male rats (56). Rats were anesthetized with ketamine
(10 mg/kg) and xylazine (5 mg/kg), and surgical occlusion of the left ante-
rior descending coronary artery was performed with a 6-0 silk suture (John-
son and Johnson). 60 minutes after occlusion, the myocardium was reper-
fused, and 3 μg of protein was injected into 3 sites from the injured region
to the border using a Hamilton syringe (Hamilton Co.) with a 30-gauge
needle. Throughout the operation, rats were ventilated with 95% O2 and
5% CO2 using a Harvard ventilator. Six animals per group were used for
morphologic analysis at 1 week after the operation. For functional studies,
we performed echocardiography at 3 weeks. To determine infarct size, the
1892?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
hearts were removed, perfused with 1% 2,3,5-triphenyltetrazolium chloride
(TTC; Sigma-Aldrich) solution, pH 7.4, at 37°C for 20 minutes, and then
fixed in 10% PBS-buffered formalin overnight at 2–8°C.
Histology and immunohistochemistry of the rat MI model. The heart was excised,
fixed with 10% PBS-buffered formalin for 24 hours, and embedded in par-
affin. Sections (5-μm thick) were mounted on gelatin-coated glass slides
to allow the use of different stains on sections. After deparaffinization and
rehydration, the tissue sections were stained with Masson’s trichrome for
analysis of fibrosis. For histological analysis, an R.T.U. VECTASTATIN
Universal Quick Kit (Vector Laboratories) was used for paraffin sections.
Sections were deparaffinized, rehydrated, and rinsed with PBS. Antigen
retrieval was performed by microwaving for 10 minutes in 10 mM sodium
citrate (pH 6.0). Sections were incubated in 3% H2O2 to quench endogenous
peroxidase activity. Sections were blocked in 2.5% normal horse serum and
incubated in primary antibody (CD31). Biotinylated pan-specific univer-
sal secondary antibody and streptavidin/peroxidase complex reagent were
used for the heart sections, which were stained with antibody using a DAB
substrate kit (Vector Laboratories). Sections were counterstained with
1% methyl green and dehydrated in 100% N-butanol, ethanol, and xylene
before mounting in VectaMount Medium (Vector Laboratories).
Tissue TUNEL assay. The TUNEL assay was performed according to the
manufacturer’s instructions (Chemicon International). A positive control
sample was prepared from a normal heart section by treating the section
with DNase I (10 U/ml, 10 minutes at room temperature). Sections were
pretreated with 3.0% H2O2, subjected to TdT enzyme at 37°C for 1 hour,
and incubated in digoxigenin-conjugated nucleotide substrate at 37°C
for 30 minutes. Nuclei exhibiting DNA fragmentation were visualized by
adding 3,3-diaminobenzidine (DAB; Vector Laboratories) for 5 minutes.
Finally, sections were counterstained with methyl green and analyzed with
light microscopy. For each group, 6 sections were prepared, and 10 differ-
ent regions were observed per section (×200).
Echocardiography. Transthoracic echocardiographic studies were performed
by an experienced cardiologist blinded to group assignments, using a GE
Vivid Seven ultrasound machine (GE Medical System) with a 10.0-MHz
transducer. Rats received general anesthesia and were placed in the left lat-
eral decubitus position. The echo transducer was placed on the left hemi-
thorax, and short axis views were recorded. Two-dimensional images were
obtained at midpapillary level (56). M-mode tracing of LV contraction was
obtained at the same level as the short-axis view. LVEDD and LVESD were
measured with M-mode tracing. Percentage of FS was calculated using the
following equation: ([LVEDD – LVESD]/LVEDD) × 100 (%). LV end diastolic
volume (LVEDV) was calculated as 7.0 × LVEDD3/(2.4 + LVEDD); LV end
systolic volume (LVESV) as 7.0 × LVESD3/(2.4 + LVESD); and LV EF as EF (%)
= (LVEDV – LVESV)/LVEDV × 100. Two images per view were obtained, and
each parameter was measured from 3 consecutive beats per image. Six values
of each parameter were obtained and averaged. Echocardiograms were stored
digitally and analyzed with EchoPAC with custom 2D strain rate imaging
software. More than 3 images were obtained in the short axis view, and the
parameters were measured from 3 consecutive beats in each image.
RT-PCR analysis. Total RNA was isolated from HUVECs using a TRIzol
reagent kit. Different amounts of total RNA (0.5–5 μg) were amplified with
RT-PCR, and the correlation between the amount of RNA used and the level
of PCR products obtained from target mRNAs and the internal standard
(GAPDH) mRNA was examined. Briefly, cDNA was synthesized from target
RNA using 200 U of reverse transcriptase and 500 ng of oligo(dT) primer in
50 mM Tris-HCl (pH 8.3), 75 mM KCl, 3 mM MgCl2, 10 mM DTT, and 1 mM
dNTPs at 42°C for 1 hour. The reaction was stopped by heating at 70°C for
15 minutes. A total of 1 μl of the cDNA mixture was then used for enzymatic
amplification. PCR was performed in 50 mM KCl, 10 mM Tris-HCl (pH 8.3),
1.5 mM MgCl2, 0.2 mM dNTPs, 2.5 U of TaqDNA polymerase (Promega),
and 0.1 μM of each primer for target genes in a DNA thermal cycler (model
PTC-200; MJ Research) under the following conditions: denaturation at
94°C for 5 minutes for the first cycle and for 30 seconds starting from the
second cycle, annealing at 55°C for 30 seconds, and extension at 72°C for
30 seconds for 25 cycles. Final extension was at 72°C for 10 minutes.
The following primer pairs were designed for RT-PCR and synthesized
by Bioneer Inc.: mDkk2_A, 5′-CAGAGATGGGATGTGTTGCC-3′ and
5′-CCTGATGGAGCACTGGTTTG-3′; mDkk2_B, 5′-GATGGGTTTT-
GTTGTGCTCG-3′ and 5′-ATGTTTCAGGTTCAGGGGGA-3′; mGapdh,
5′-CGCCACAGTTTCCCGGAGGG-3′ and 5′-CCCTCCAAAATCAAGT-
GGGG-3′; DKK1, 5′-CCTGGAGTGTAAGAGCTTTG-3′ and 5′-CCAAGA-
GATCCTTGCGTT-3′; DKK2, 5′-TAGAGATTGAGTTTGAGCCT-3′ and
5′-AAAGGGTGGACATAAGAAA-3′; KREMEN2, 5′-CACCGACTGTGAC-
CAGAT-3′ and 5′-GAGTAGATGACGCCCTGAG-3′; LRP6, 5′-GTCCTTC-
CACTCATAGGTCA-3′ and 5′-GTGGGTAGAGGTGATGAGAA-3′; LRP5,
5′-GGGTGGTGTCTATTTTGTGT-3′ and 5′-CCGGAATGTTTGAAGAG-
TAG-3′; APC, 5′-CAGGCAAAACAAAATGTGGG-3′ and 5′-CGCTTAG-
GACTTTGGGTTCC-3′; Asef2, 5′-TCTACTCGGGGGAGCTGACT-3′ and
5′-CCTGCTTCCTCTGCAGTTCA-3′; GAPDH, 5′-CGCCACAGTTTCCC-
GGAGGG-3′ and 5′-CCCTCCAAAATCAAGTGGGG-3′.
Cdc42 assay. Cdc42 activity was measured with a CDC42 activation kit
(Upstate Biotechnology) according to the manufacturer’s instructions.
HUVECs were washed once with ice-cold PBS and lysed with lysis buf-
fer containing 25 mM HEPES, pH 7.5, 150 mM NaCl, 1% Igepal CA-630,
10 mM MgCl2, 1 mM EDTA, 10% glycerol, 1 mM Na3VO4, 10 μg/ml apro-
tinin, 10 μg/ml leupeptin, and 25 mM NaF for 15 minutes at 4°C. Insoluble
materials were removed by centrifugation. Five micrograms of PAK1-agarose
beads, which specifically bind to active Cdc42, were added to the cell lysates
and incubated for 1 hour at 4°C. Agarose beads were washed with lysis buf-
fer 3 times and boiled in 2× Laemmli sample buffer. Samples were resolved
by SDS-PAGE and immunoblotted with an anti-Cdc42 antibody.
Animal studies. All mice were maintained in a laminar airflow cabinet
under specific pathogen–free conditions. All facilities were approved by the
AAALAC (Association of Assessment and Accreditation of Laboratory Ani-
mal Care). All animal experiments were conducted under the institutional
guidelines established for the Animal Core Facility at Yonsei University
College of Medicine and were approved by the IACUC, Yonsei University.
Statistics. Data are presented as mean ± SD or ± SEM. Statistical com-
parisons between groups were performed using 1-way analysis of variance
(ANOVA) followed by Student’s t test (1-tailed).
This work was supported by a National Research Laboratory grant
(ROA-2007-000-20099-0); a Korea Biotech R&D Group grant
(F104AC010003-07A0301) from the Korean Science and Engi-
neering Foundation (KOSEF) funded by the Ministry of Educa-
tion & Science Technology; a grant from the Korea Health 21 R&D
Project, Ministry of Health Welfare and Family Affairs, Republic of
Korea (A085136); a Korea Research Foundation grant funded by
the Korean government (KRF-2006-351-C00021 to J.K. Min); and
a grant from the Korea Research Council of Fundamental Science
& Technology (Stem Cell Research Program to J.K. Min).
Received for publication February 4, 2010, and accepted in revised
form February 16, 2011.
Address correspondence to: Young-Guen Kwon, Department of Bio-
chemistry, College of Life Science and Biotechnology, Yonsei Uni-
versity, Seoul, 120-749, Republic of Korea. Phone: 82.2.2123.5697;
Fax: 82.2.362.9897; E-mail: firstname.lastname@example.org.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 5 May 2011
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