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RESEARCH ARTICLE 4133
DEVELOPMENT AND STEM CELLS
Development 139, 4133-4142 (2012) doi:10.1242/dev.079756
© 2012. Published by The Company of Biologists Ltd
Migration of cardiomyocytes is essential for heart
regeneration in zebrafish
Junji Itou1,2, Isao Oishi3, Hiroko Kawakami1,2, Tiffany J. Glass4, Jenna Richter1, Austin Johnson1,
Troy C. Lund4, and Yasuhiko Kawakami1,2,5,*
SUMMARY
Adult zebrafish possess a significant ability to regenerate injured heart tissue through proliferation of pre-existing cardiomyocytes,
which contrasts with the inability of mammals to do so after the immediate postnatal period. Zebrafish therefore provide a model
system in which to study how an injured heart can be repaired. However, it remains unknown what important processes
cardiomyocytes are involved in other than partial de-differentiation and proliferation. Here we show that migration of
cardiomyocytes to the injury site is essential for heart regeneration. Ventricular amputation induced expression of cxcl12a and cxcr4b,
genes encoding a chemokine ligand and its receptor. We found that cxcl12a was expressed in the epicardial tissue and that Cxcr4
was expressed in cardiomyocytes. We show that pharmacological blocking of Cxcr4 function as well as genetic loss of cxcr4b function
causes failure to regenerate the heart after ventricular resection. Cardiomyocyte proliferation was not affected but a large portion
of proliferating cardiomyocytes remained localized outside the injury site. A photoconvertible fluorescent reporter-based
cardiomyocyte-tracing assay demonstrates that cardiomyocytes migrated into the injury site in control hearts but that migration
was inhibited in the Cxcr4-blocked hearts. By contrast, the epicardial cells and vascular endothelial cells were not affected by blocking
Cxcr4 function. Our data show that the migration of cardiomyocytes into the injury site is regulated independently of proliferation,
and that coordination of both processes is necessary for heart regeneration.
INTRODUCTION
Regeneration is a complex biological process by which animals
restore the shape, structure and function of body parts lost to injury
or experiment (Brockes and Kumar, 2005; Poss, 2010). One of the
most remarkable examples is heart regeneration (Ausoni and
Sartore, 2009; Laflamme and Murry, 2011). Several nonmammalian vertebrates, including zebrafish, possess the significant
ability to restore injured heart. After resectioning of the ventricular
apex, zebrafish restore lost heart tissue within 1-2 months (Poss et
al., 2002; Raya et al., 2003), which contrasts to the inability of
mammals to do so after the immediate postnatal period (Bergmann
et al., 2009; Porrello et al., 2011). The regenerative ability is also
observed in response to a variety of injuries, including cryoprobeinduced injury (Chablais et al., 2011; González-Rosa et al., 2011;
Schnabel et al., 2011) and genetic ablation of cardiomyocytes
(CMs) by transgenic induction of toxin expression (Wang et al.,
2011). It therefore provides a model system to study how an injured
heart can be repaired (Poss, 2007; Raya et al., 2004). Recent
reports showed that the major source of regenerated CMs was preexisting CMs, rather than stem/progenitor cells, that undergo partial
de-differentiation, re-enter the cell cycle and proliferate to restore
the injured heart (Jopling et al., 2010; Kikuchi et al., 2010).
1
Department of Genetics, Cell Biology and Development, University of Minnesota,
Minneapolis, MN 55455, USA. 2Stem Cell Institute, University of Minnesota,
Minneapolis, MN 55455, USA. 3Health Research Institute, National Institute of
Advanced Industrial Science and Technology, Ikeda, Osaka 563-8577, Japan.
4
Division of Pediatric Blood and Marrow Transplant, University of Minnesota,
Minneapolis, MN 55455, USA. 5Developmental Biology Center, University of
Minnesota, Minneapolis, MN 55455, USA.
*Author for correspondence ([email protected])
Accepted 18 August 2012
Simultaneously, vascularization of the regenerating area takes
place, which requires the activities of fibroblast growth factor
(FGF) signaling and vascular endothelial growth factor (VEGF)
signaling (Kim et al., 2010; Lepilina et al., 2006). However, it is
still unknown whether CMs are involved in other processes, in
addition to proliferation, to restore the injured heart. More
specifically, cell migration is known to be involved in the
development of a variety of organs (Friedl and Gilmour, 2009; Raz
and Mahabaleshwar, 2009), and organ regeneration involves
reactivation of developmental processes and genes (Iovine, 2007).
The recent advent of photoconvertible fluorescent reporter
proteins provided a new approach for cell migration analysis (Stark
and Kulesa, 2007). Natural and engineered green fluorescent
proteins, which can be photoconverted into red fluorescent
proteins, have been developed (Lukyanov et al., 2005). Such
proteins include Kaede (Ando et al., 2002), PA-GFP (Patterson and
Lippincott-Schwartz, 2002), KikGR (Tsutsui et al., 2005), EosEP
(Nienhaus et al., 2006), PA-mRFP (Verkhusha and Sorkin, 2005)
and Dendra (Gurskaya et al., 2006). Kaede is one of the founding
members of photoconvertible fluorescent proteins, serendipitously
found from a stony coral, Trachyphyllia geoffroyi (Ando et al.,
2002). The unique photoconvertible characters of Kaede represent
several advantages for optic marking. First, the red state is
comparable to the green in terms of brightness and stability.
Second, photoconversion is shown to result in a more than 2000fold increase in the red/green fluorescent ratio. Third, completely
separate wavelengths of light can be used for observation and
photoconversion. By taking advantage of these unique properties,
several reports have demonstrated cell tracing in vivo, such as
monitoring cell migration during zebrafish embryogenesis (Hatta
et al., 2006), cell migratory behaviors in cortical slices of mice
(Mutoh et al., 2006) and T-cell migration during cutaneous immune
response in mice (Tomura et al., 2010; Tomura et al., 2008). The
DEVELOPMENT
KEY WORDS: Heart regeneration, Cardiomyocytes, Zebrafish, Directed migration, CXCL12-CXCR4
Development 139 (22)
4134 RESEARCH ARTICLE
MATERIALS AND METHODS
Zebrafish maintenance and surgery
Zebrafish were maintained under standard conditions at around 28°C, and
adult zebrafish (6 to 18 months old) were used for experiments. Ventricular
amputation was performed as previously published (Raya et al., 2003).
Care and experimentation were done in accordance with the Institutional
Animal Care and Use Committee of the University of Minnesota and the
National Institute of Advanced Industrial Science and Technology, Japan.
In situ hybridization, immunostaining and TUNEL assay
In situ hybridization on sections 14 m thick was performed as previously
described (Kawakami et al., 2011). Immunostaining on sections was
performed according to a standard procedure (Kawakami et al., 2006; Raya
et al., 2003). The primary antibodies used were anti-myosin heavy chain
(MHC; Developmental Studies Hybridoma Bank, MF20, 5.14 g/ml), antiCXCR4 (Santa Cruz Biotechnology, sc-6190, 1:100), anti-PCNA (Santa
Cruz Biotechnology; sc-56, 1:100), anti-MEF2 (Santa Cruz Biotechnology;
sc-313, 1:50), anti-GFP (Molecular Probes, A11122, 1:500) and antiDsRed2 (Clontech, 632496, 1:500). Secondary antibodies used were Alexa
fluorophore-labeled anti-mouse or rabbit or goat IgG (Invitrogen, 1:1000)
or biotinylated anti-mouse IgG (Vector Laboratories, BA-9200, 1:500). An
In Situ Cell Death Detection Kit (Roche Diagnostics) was used for TUNEL
assay according to the manufacturer’s instruction. Counterstaining was
done with DAPI or hematoxylin. The number of proliferating CMs was
counted manually with the sections that represent the largest injury area at
the center of the injury site of each heart.
EdU-labeling experiment
To label proliferating cells, 20 l of 0.5 mg/ml EdU solution was
intraperitoneally injected into zebrafish at 7 days post amputation (dpa) and
10 dpa. Hearts were collected at 13 dpa and subjected to standard analysis
according to the manufacturer’s instructions (Invitrogen, Click-IT EdU cell
proliferation assays).
Pharmacological blocking of Cxcr4 function and genetic loss of
cxcr4b function
For pharmacological blocking of Cxcr4 function, each fish was maintained
in 50 ml system water with or without 40 nM FC131 after surgery (Narumi
et al., 2010) (Wako Chemical, Osaka, Japan), and the water was refreshed
daily. Because the carrier is H2O, nothing was added into the containers of
control fish. To assist with the penetration of FC131, the pericardiac cavity
was surgically opened weekly in both control and treated fish. For genetic
loss of cxcr4b function, we used the odysseus mutant fish line, which
possesses a null mutation in the cxcr4b gene (Knaut et al., 2003). Wildtype siblings were used as controls for the odysseus mutants (hereafter
odysseus mutants are referred to as cxcr4b–/–).
Generation of the cmlc2a-Kaede line
The cmlc2a-Kaede line was established with the zebrafish cmlc2a promoter
(Huang et al., 2003) and Kaede (Medical & Biological Laboratories,
Nagoya, Japan) using the Tol2 transposon system (Kawakami et al., 2004;
Urasaki et al., 2006). The P0 fish were outcrossed with AB fish, and stable
transgenic lines were established after evaluating cardiac-specific
expression of Kaede.
Photoconversion of Kaede
For photoconversion of cmlc2a-Kaede transgenic fish hearts, we made a
small window by manually dissecting the pericardiac cavity after
anesthetizing fish. A compound microscope (Nikon LABPHOT) equipped
with a 100 watt mercury lamp, diachronic mirror of 380 nm and a barrier
filter of 420 nm were used to irradiate the heart for 90 seconds with
approximately 2 mm distance between the 20⫻ objective lens and the
heart. To injure the heart, we enlarged the pericardiac window and
amputated the apex of the ventricle under fluorescent monitoring with a
Zeiss V12 stereomicroscope.
Imaging
Fluorescent confocal images were obtained by using a Zeiss LSM 710 laser
scanning microscope system, and analyzed by ZEN2009 software. For
Kaede imaging, non-fixed hearts were embedded in optimal cutting
temperature compound and frozen. Sections at 20 m thickness were dried,
washed with PBS, rinsed with water and mounted for confocal imaging.
Kaede green fluorescence was excited by 514 nm light and 519–556 nm
fluorescence was detected. Wavelengths for excitation and detection of
Kaede red fluorescence were 562 nm and 566–674 nm, respectively.
Statistical analysis
Statistical significance was analyzed by the Student’s t-test, and shown as
average ± standard deviation. P-values are indicated within each panel.
RESULTS
Induction of cxcl12a and cxcr4b expression after
heart injury
During expression screening of genes expressed in developing and
regenerating zebrafish hearts, we found that cxcl12a, a gene
encoding a chemokine ligand (also known as sdf1a), and its
receptor cxcr4b were expressed during heart regeneration (Fig. 1).
The transcripts of cxcl12a were detected between 3 dpa and 7 dpa
at the surface and inside of the regenerating area (Fig. 1). The
signals of cxcr4b became evident at 5 dpa, and persisted until 10
dpa (Fig. 1). cxcr4b expression was detected at the surface layer of
the regenerating area and around the injury site. These expression
patterns were not detected in sham-operated hearts at 3 dpa
(cxcl12a) and 7 dpa (cxcr4b), the stages at which they were
detected at high levels (n5 each, data not shown). By contrast, we
did not detect transcripts of the related ligand-receptor pair cxcl12b
and cxcr4a during heart regeneration by in situ hybridization
(supplementary material Fig. S1). The CXCL12-CXCR4 system is
known to regulate directed cell migration during embryonic
development (Friedl and Gilmour, 2009; Raz and Mahabaleshwar,
2009; Schier, 2003). Thus, induction of the expression of cxcl12a
and cxcr4b after heart injury suggests that directed cell migration
plays a role during heart regeneration.
CMs express cxcr4b
As a first step to investigate the possible involvement of directed
cell migration during heart regeneration, we sought to identify cell
types that express cxcr4b and cxcl12a. Immunofluorescence
analysis showed that the CM-specific nuclear signal of Mef2
(supplementary material Fig. S2) is associated with membranelocalized Cxcr4 signal on the same confocal plane (Fig. 2A,B),
indicating that CMs express Cxcr4b. Further analysis of the Cxcr4
signal and CM-specific cmlc2a-EGFP signal also showed that 287
Cxcr4-positive signals from six sections were associated with the
cmlc2a-EGFP signal (Fig. 2C-E). Similarly, the Cxcr4 signal was
DEVELOPMENT
use of cell-type-specific promoters/enhancers has also facilitated
monitoring specific types of cells in combination with
photoconversion technology, such as differentiation of
cardiomyocytes (de Pater et al., 2009), beta cells of the pancreas
(Pisharath et al., 2007) and postmitotic neurons (Pan et al., 2012).
Here we show that migration of CMs to the injury site is essential
for heart regeneration. We found that the chemokine ligand, cxcl12a
and its receptor, cxcr4b, were induced after heart injury in zebrafish.
Blocking Cxcr4 function caused mis-localization of proliferating
CMs outside of the injury site without affecting their proliferation.
By localized photoconversion in hearts of the CM-specific Kaede
transgenic zebrafish, we show that CMs migrate toward the injury
site only after heart damage, and this migration requires Cxcr4
function. Our data show that the migration of CMs into the injury
site is regulated independently of proliferation, and that coordination
of both processes is essential for heart regeneration.
Fig. 1. Expression of cxcl12a and cxcr4b during heart
regeneration. (A-N)In situ hybridization for cxcl12a (A,C,E,G,I,K,M)
and cxcr4b (B,D,F,H,J,L,N) in the regenerating heart. Time points at 0
(A,B), 1 (C,D), 3 (E,F), 5 (G,H), 7 (I,J), 10 (K,L) and 14 (M,N) dpa are
shown. An evident expression of cxcl12a was detected at 3 dpa to 7
dpa (E,G,I). Strong expression of cxcr4b was detected at 5 dpa (H) and
persisted until 10 dpa (L). Scale bar: 50m.
also associated with MHC and Mef2 signals (Fig. 3). Moreover,
optic sectioning of Cxcr4-expressing CMs showed Cxcr4 signal in
the cytoplasm, in addition to the membrane (Fig. 3B,C), suggesting
Cxcl12a-dependent internalization of Cxcr4b (Orsini et al., 1999).
These results demonstrate that CMs express Cxcr4. In the mouse,
endothelial cells also express Cxcr4 (Gupta et al., 1998; Volin et
al., 1998); however, we did not detect Cxcr4 in endothelial cells
(Fig. 2F-H). Two hundred and sixty Cxcr4 signals from five
sections of fli1-EGFP transgenic fish heart were not associated
with the endothelial-cell-specific fli1-EGFP signal.
Next, we investigated which cells express cxcl12a. Consistent
with the expression of cxcl12a at the surface of the regenerating
area (Fig. 1), the cxcl12a signals overlapped with wt1b signals
(supplementary material Fig. S3A-C⬘), which is expressed in the
epicardial cells (Perner et al., 2007). We did not detect expression
of the cxcl12a-DsRed2 reporter (Glass et al., 2011) in MHCpositive CMs (supplementary material Fig. S3D-F) or in fli1EGFP-positive endothelial cells (supplementary material Fig. S3GI). These data indicate that epicardial cells express cxcl12a after
heart injury.
Cxcr4 function is required for heart regeneration
To determine the functional significance of the Cxcl12-Cxcr4
system during heart regeneration, we blocked Cxcr4 function by
treating fish with an antagonist after ventricular amputation. We
first tested whether FC131, a recently developed selective CXCR4
antagonist (Narumi et al., 2010), can block zebrafish Cxcr4
RESEARCH ARTICLE 4135
Fig. 2. Cxcr4 is present on CMs. (A,B)Confocal images of Cxcr4
(green) and Mef2 (magenta) double staining of control heart at 7 dpa.
The Cxcr4 signal was detected at the cell surface of nuclear Mef2positive cells in the regenerating area (arrowheads in B). B shows a
higher magnification image of the boxed area in A. The dotted line
indicates the amputation plane. (C-E)Confocal images of Cxcr4 (C) and
cmlc2a-EGFP signal, detected by anti-GFP antibody (D), and a merged
image (E). The arrowheads point to cells positive for both Cxcr4 and
cmlc2a-EGFP signals. (F-H)Confocal images of Cxcr4 (F) and fli1-EGFP
signal, detected by anti-GFP antibody (G), and a merged image (H).
Scale bars: 50m.
function. AMD3100, a widely used CXCR4 antagonist, was not
used because our previous studies indicated that it did not
efficiently block Cxcr4 function, either in the adult zebrafish
hematopoietic system in vivo (Glass et al., 2011) or in the
migration of primordial germ cells in zebrafish embryos (data not
shown). Treating zebrafish embryos with FC131 significantly
impaired directed migration of primordial germ cells into the
gonad-forming region (supplementary material Fig. S4), a process
that has been shown to depend on cxcr4b function (Doitsidou et al.,
2002; Knaut et al., 2003), demonstrating that FC131 can effectively
block Cxcr4 function in zebrafish. We then treated zebrafish with
FC131 after amputation of the ventricle and found that this resulted
in failure to regenerate the heart (Fig. 4A-F⬘). In control
regenerating hearts, MHC signals (CM marker) were strongly
detected in the injury site, where the heart tissue was removed, at
14 dpa, and the regenerating tissue was filled with CMs at 30 dpa
(Fig. 4A,A⬘,C,C⬘). At 60 dpa, the regenerated myocardium
exhibited a strong MHC signal, comparable to the non-injured area
(Fig. 4E,E⬘). However, significantly lower levels of MHC signals
were detected in the injury site of CXCR4-antagonist-treated hearts
at 14 dpa (Fig. 4B,B⬘, n4/5 hearts). After 30 days and 60 days the
CXCR4-antagonist-treated heart lacked a sealed wall (Fig. 4D,D⬘,
red arrows, n5/5 hearts; 4F,F⬘, n3/4 hearts), which was fully
formed in control hearts (Fig. 4C,C⬘, black arrows, n5/5 hearts;
4E,E⬘, n5/5 hearts). These results indicate that the CXCR4antagonist-treated heart did not regenerate. The significant
reduction of MHC signals in the injury site was also observed in
cxcr4b–/– fish at 14 dpa (Fig. 4G,H⬘, n3/3 hearts). Similar to the
CXCR4-antagonist-treated heart, cxcr4b–/– fish hearts failed to
form the sealed wall at 30 dpa (Fig. 4J,J⬘, n2/2 hearts) and 60 dpa
(Fig. 4L,L⬘, n2/2 hearts), compared with control hearts (Fig. 4I,I⬘,
DEVELOPMENT
Cardiomyocyte migration
4136 RESEARCH ARTICLE
Development 139 (22)
Fig. 3. Confocal imaging of Cxcr4-expressing cardiomyocytes.
(A)Confocal images of MHC, Cxcr4 and Mef2 at 7 dpa. The yellow
arrowheads point to Cxcr4-positive cells. (B)Images of a Cxcr4expressing CM at 500 nm distance along the z-axis. Mef2 represents
the nucleus (magenta), MHC represents cytoplasm (green) and Cxcr4 is
shown in yellow. The numbers in the upper area of each panel indicate
positions along the z-axis. (C)Optical section of a Cxcr4-expressing cell.
High levels of Mef2 signal and MHC signal define a nucleus and
cytoplasm, respectively. The open arrowheads and solid arrowheads
indicate Cxcr4 in membrane and cytoplasm, respectively. The vertical
axis represents fluorescence intensity. Scale bars: 100m in A; 10m
in B.
n3/3 hearts; 4K,K⬘, n3/3 hearts). Fibrin clearance at 14 dpa was
similar in control and cxcr4–/– fish (supplementary material Fig. S5,
n2/2 for both control and cxcr4–/– hearts), and thus, the failure to
regenerate the heart is unlikely to be caused by abnormal fibrin
deposition. These data demonstrate that cxcr4b function is required
for heart regeneration in zebrafish. These results strongly support
our hypothesis that cxcr4b functions in CMs that contribute to heart
regeneration. Although cxcr4b expression was specific to CMs in
regenerating zebrafish hearts (Fig. 2), Cxcr4 is also known to be
involved in heart development in mice (Ma et al., 1998; Tachibana
et al., 1998; Zou et al., 1998). In order to avoid complications
caused by an unidentified role of cxcr4b in zebrafish heart
development, if any, we focused our study of cxcr4b function by
pharmacological antagonism using FC131.
Mis-localization of proliferating CMs by blocking
Cxcr4 function
Given that the proliferation of CMs is a major factor for heart
regeneration (Jopling et al., 2010; Kikuchi et al., 2010), we asked
whether blocking Cxcr4 function affected CM proliferation and/or
survival. The number of proliferating CMs, visualized by
proliferating cell nuclear antigen (Pcna, an S-phase marker) and
Mef2 (CM marker), did not change significantly in Cxcr4-blocked
hearts compared with control hearts at 7, 14, 21 and 30 dpa (Fig.
5A-C, n5 hearts at each time point). This result also indicates that
the failure to regenerate the heart by blocking Cxcr4 function is not
due to delayed CM proliferation. The number of Pcna-positive
cells, which includes CMs and non-CMs, was also not changed at
these time points (supplementary material Fig. S6), indicating that
the proliferation of other cell types was also unaffected by CXCR4
antagonist treatment. TUNEL analysis did not show an increase in
the number of cell deaths at 7 dpa and 14 dpa (supplementary
material Fig. S7). These data indicate that the failure to regenerate
the amputated heart by blocking Cxcr4 function is not caused by
defects in proliferation or survival of CMs and other cells. A recent
study showed that the number of genetically labeled CMs located
at the sub-epicardial area increases in the injury site, whereas their
number reduces at the edge of the injury site during heart
regeneration (Kikuchi et al., 2010). Thus, we compared the number
of proliferating CMs in the injury site, similar to the study by
Kikuchi and colleagues (Kikuchi et al., 2010). We found a change
in the localization of proliferating CMs in Cxcr4-blocked hearts. In
control hearts we found 74.0% of proliferating CMs around the
center of the regenerating area, and the majority of the other
proliferating CMs at the edge of the regenerating area at 14 dpa.
By contrast, only 23.5% of the proliferating CMs were located
around the center of the injury site in Cxcr4-blocked hearts (Fig.
5D). Because the total number of proliferating CMs was not altered
(Fig. 5C), this observation suggests that Cxcr4 function is
necessary for localization of proliferating CMs in the regenerating
area.
Detection of Pcna visualizes cells in the S phase of the cell cycle
at the time of sample fixation. In order to further examine CM
DEVELOPMENT
Fig. 4. Cxcr4 function is required for heart regeneration.
(A-F⬘) MHC (brown) and hematoxylin (purple) staining of control
(A,A⬘,C,C⬘,E,E⬘) and CXCR4-antagonist-treated (B,B⬘,D,D⬘,F,F⬘) heart at
14 dpa (A-B⬘), 30 dpa (C-D⬘) and 60 dpa (E-F⬘). (G-L⬘) MHC and
hematoxylin staining of wild-type (G,G⬘,I,I⬘,K,K⬘) and cxcr4b–/–
(H,H⬘,J,J⬘,L,L⬘) heart at 14 dpa (G-H⬘), 30 dpa (I-J⬘) and 60 dpa (K-L⬘).
A⬘-L⬘ show close up of the boxed area in A-L, respectively. The
arrowheads in A⬘, B⬘, G⬘ and H⬘ point to the MHC signal in the
regenerating area. The black and red arrows in C⬘-F⬘ and I⬘-L⬘ point to
the presence and absence of a sealed muscle wall, respectively. Dotted
lines in A, B, G and H indicate the amputation planes. Scale bars:
50m.
Cardiomyocyte migration
RESEARCH ARTICLE 4137
experiment further confirmed a role for Cxcr4 function in the
localization of proliferating CMs in the regenerating area.
proliferation, we labeled cells by EdU. Injection of EdU at 7 dpa
and 10 dpa, followed by detection at 13 dpa, allows for labeling
cells that underwent cell division during 7 dpa to 13 dpa (Fig. 5EF⬘). Similar to the analysis of Pcna detection, the number of
proliferating CMs, visualized as EdU and cmlc2a-mCherry doublepositive cells, were at similar levels in control and CXCR4antagonist-treated hearts (Fig. 5G). We found that 72.4% of EdUlabeled CMs were located around the center of the regenerating
area. By contrast, only 12.4% of EdU-labeled CMs were located
around the center of the injury site in Cxcr4-blocked hearts (Fig.
5H), similar to the case with Pcna analysis. This EdU-labeling
Cxcr4-dependent CM migration after injury
To analyze the migration of CMs, we amputated the apex in the
non-irradiated portions of the ventricle under a fluorescent
stereomicroscope (Fig. 7A-C, n6). Three days after amputation,
we did not detect CMs in the regenerating area (data not shown,
n7), consistent with the lack of cxcr4b signal in the regenerating
area at this time point (Fig. 1F). However, seven days after
amputation, CMs with both green and red fluorescence, and CMs
with only green fluorescence, were detected in the regenerating
area (Fig. 7D,E, n6). We detected 5.5% of photoconverted CMs
in the regenerating area (Fig. 7P). This result demonstrates that
CMs migrated into the regenerating area by 7 dpa. The ratio of
photoconverted CMs in the regenerating area was 10.4% and
12.1% at 10 dpa (Fig. 7H,I,P, n7 hearts) and 14 dpa (Fig. 7L,M,P,
n7 hearts), respectively. At 10 dpa and 14 dpa, a portion of
photoconverted CMs exhibited strong green signals due to
continued de novo production of Kaede. Blocking Cxcr4 function
caused CMs to be excluded from the injury site at 7 dpa (Fig.
7F,G,P, n6 hearts), 10 dpa (Fig. 7J,K,P, n6 hearts) and 14 dpa
(Fig. 7N,O,P, n6 hearts). Our data revealed that the cxcl12acxcr4b-dependent system regulates directed migration of CMs into
the injury site, and that this process is essential for heart
regeneration.
DEVELOPMENT
Fig. 5. Blocking Cxcr4 function causes mis-localization of
proliferating CMs. (A-B⬘) Pcna (green) and Mef2 (magenta) staining
of control (A,A⬘) and CXCR4-antagonist-treated (B,B⬘) hearts at 14 dpa.
The yellow arrowheads point to proliferating CMs seen as white signal.
A⬘ and B⬘ show close up of the boxed area in A and B. For simplicity,
not all proliferating CMs are labeled. (C)Quantitation of the number of
proliferating CMs in control and CXCR4-antagonist-treated hearts at 7,
14, 21 and 30 dpa. The vertical axis represents the number of
proliferating CMs per section. A section that represents the largest
injury area at the center of the injury site of each heart was examined
(n5). P-values at each time point are shown. (D)Quantitation of the
ratio of proliferating CMs in the injury site compared with the number
of entire proliferating CMs in control and CXCR4 antagonist-treated
hearts at 14 dpa. The same samples examined in C were used.
(E-F⬘) EdU (green) and cmlc2a-mCherry (magenta) signal of control
(E,E⬘) and CXCR4 antagonist-treated (F,F⬘) hearts at 13 dpa. The yellow
arrowheads point to proliferating CMs seen as white signal. For
simplicity, not all proliferating CMs are labeled. E⬘ and F⬘ show close up
of the boxed area in E and F. (G)Quantitation of the number of
EdU/cmlc2a-mCherry double-positive cells in control and CXCR4
antagonist-treated hearts at 13 dpa. Vertical axis represents the number
of proliferating CMs per section. A section that represents the largest
injury area at the center of the injury site of each heart was examined
(n5). (H)Quantitation of the ratio of EdU-positive CMs in the injury
site compared with the number of entire EdU-positive CMs in control
and CXCR4-antagonist-treated hearts at 13 dpa. The same samples
examined in G were used. For D, G and H, P-values by Student’s t-test
are shown. Dotted lines in A, B, E and F indicate the amputation
planes. Scale bars: 50m.
Localized labeling of CMs by photoconversion of
the Kaede fluorescent protein
Given that the Cxcl12-Cxcr4 system is known to regulate directed
cell migration in a variety of processes (Friedl and Gilmour, 2009;
Raz and Mahabaleshwar, 2009; Schier, 2003), the abnormal
localization of proliferating CMs suggests that the migration of
proliferating CMs, rather than local activation of CMs in the injury
site, is necessary for heart regeneration. To address this hypothesis,
we developed a fluorescent reporter-based CM-tracing assay. We
established a transgenic zebrafish line that expresses Kaede (Ando
et al., 2002), driven by the CM-specific cmlc2a promoter
(supplementary material Fig. S8A,B, cmlc2a-Kaede). The green
fluorescence of Kaede can be irreversibly converted into stable red
fluorescence by ultraviolet irradiation for cell tracing in vivo
(supplementary material Fig. S8C-H) (Ando et al., 2002; Tomura
et al., 2008). Thus, we were able to label CMs in a restricted area
of the ventricle with red fluorescence in order to trace labeled CMs
during heart regeneration (Fig. 6A-F; Fig. 7A). Maintaining
cmlc2a-Kaede fish under normal breeding conditions (Fig. 6G-J,
n8), as well as ventricular amputation under a fluorescent
stereomicroscope (Fig. 6K-N, n8), did not induce green to red
photoconversion. For localized photoconversion of the heart, we
exposed a small part of the ventricle and performed irradiation (see
Materials and methods) (Fig. 6A-F). Photoconverted red
fluorescence was observed immediately after irradiation and only
in the irradiated area (Fig. 6O-R, n8). Seven days after irradiation,
the red fluorescence was stable and remained as a cluster (Fig. 6SV, n8), indicating that CMs labeled with red fluorescence did not
migrate under normal conditions. We also observed newly
synthesized green Kaede 7 days after irradiation such that the
irradiated CMs fluoresced both red and green (compare Fig. 6O
and 6S). This indicates that the irradiation was not toxic to CMs.
Also, we did not detect cell death at 1 day and 3 days after
irradiation in the irradiated area (data not shown).
Epicardial cells are unlikely to be affected by
blocking Cxcr4
Cells other than CMs, such as epicardial cells and vascular
endothelial cells, are also known to participate in heart
regeneration in zebrafish (Kikuchi et al., 2011b; Kim et al.,
2010; Lepilina et al., 2006). To further understand how Cxcr4
regulates heart regeneration, we examined whether these cells
are also affected by blocking Cxcr4 function during heart
regeneration. Expression of aldehyde dehydrogenase 1a2
(aldh1a2, also known as retinaldehyde dehydrogenase 2,
raldh2), a gene encoding a rate-limiting enzyme for retinoic acid
synthesis, is upregulated in the epicardial tissue after cardiac
damage (Lepilina et al., 2006), and retinoic acid signaling is
required for heart regeneration (Kikuchi et al., 2011b). We
observed aldh1a2 expression in a wide region of the epicardium
at 3 dpa, and at the surface of the injury site at 7 dpa, similar to
the control heart (supplementary material Fig. S9A-F). aldh1a2
expression is also detected in endocardial cells after heart injury
(Kikuchi et al., 2011b), which can be visualized by fli1-EGFP
reporter at 3 dpa (supplementary material Fig. S9C,D). Thus,
aldh1a2 expression in both epicardial and endocardial tissue
appeared to be unaffected by CXCR4 antagonist treatment.
Expression of wt1b marks the epicardial tissue in regenerating
hearts (González-Rosa et al., 2011; Kikuchi et al., 2011a;
Schnabel et al., 2011). We detected comparable wt1b mRNA
expression in the epicardial tissue of both control and Cxcr4blocked hearts at 3 dpa (supplementary material Fig. S9G-H⬘)
and 7 dpa (supplementary material Fig. S9I-J⬘). To further
evaluate whether epicardial responses are affected by CXCR4
antagonist treatment, we measured the length of the wt1bexpressing domain (supplementary material Fig. S9G,H,I,J, blue
lines). The ratio of the lengths of the wt1b-expressing domain to
the length of the surface of the regenerating area were
comparable in control and CXCR4-antagonist-treated hearts at
both time points (supplementary material Fig. S9K). These
results indicate that gene expression in the epicardial tissue was
not affected by blocking Cxcr4 function during heart
regeneration.
Blocking of CXCR4 function is unlikely to affect
neo-vascularization in regenerating heart
Neo-vascularization of the regenerating area is crucial for heart
regeneration (Kim et al., 2010; Lepilina et al., 2006). Although
Cxcr4 was not detected in endothelial cells after ventricular
amputation (Fig. 2), we sought to clarify whether vascularization
was affected by blocking Cxcr4 function. The fli1-EGFP signal
was detected similarly in control and CXCR4-antagonist-treated
hearts at 7 dpa (Fig. 8A,B, n3) and 14 dpa (Fig. 8C,D, n3).
Analysis of the fli1-EGFP-positive region in the regenerating area
at 14 dpa by ImageJ software showed a similar level of
vascularization (Fig. 8E). Thus, CXCR4 antagonist-treatment is
unlikely to affect neo-vascularization during heart regeneration in
zebrafish.
It has been shown that FGF signaling is required for neovascularization of the regenerating area during heart regeneration
(Lepilina et al., 2006); thus, we also examined activation of FGF
signaling. Phosphorylation of ERK, a hallmark of the activation of
FGF signaling, was detected both in control and Cxcr4-blocked
hearts (Fig. 8F,G). Expression of mkp3/dusp6, a target of FGF
signaling (Kawakami et al., 2003), was also detected similarly in
the control and CXCR4-antagonist-treated heart (Fig. 8H,I). These
results indicate that activation of FGF signaling occurred similarly
Development 139 (22)
Fig. 6. CM labeling by the Kaede photoconversion shows no CM
migration in non-injured hearts. (A-F)Localized photoconversion of
the adult cmlc2a-Kaede heart. Green (A,D), red (B,E) and merged
images (C,F) of whole mount samples without irradiation (A-C) and
immediately after irradiation (D-F) are shown. Without irradiation,
hearts show only green fluorescence (A-C). After localized irradiation,
the fluorescence was converted to red in the irradiated area (E,F). (GV)Green (G,K,O,S), red (H,L,P,T) and merged images (I,M,Q,U) of
sectioned samples are shown. J, N, R and V show higher magnification
images of the boxed area in I, M, Q and U, respectively. (G-J)No red
fluorescence was detected without irradiation and injury. (K-N)No red
fluorescence was detected at 7 dpa without irradiation. The dotted line
indicates the amputation plane. Some green cells are detected in the
regenerating area (N, open arrowheads). (O-R)Images immediately
after irradiation without injury. The green signal in the irradiated area
was lost (arrowheads, O) and red signal in the same area was detected
(arrowheads, P). The merged images show boundary of green-red
signal (Q,R). The yellow arrows in R point to the photoconverted CMs.
(S-V)Images 7 days after irradiation without injury. Red signal was
detected in the irradiated area (arrowheads, T), and green signal was
detected in the irradiated area by newly synthesized Kaede
(arrowheads, S). The yellow arrows in V point to the photoconverted
CMs. Scale bars: 500m in A; 50m in G and J.
in both control and Cxcr4-blocked hearts. Comparable activation
of gene expression in the epicardial tissue (supplementary material
Fig. S9) and vascularization (Fig. 8) further support the idea that
the failure to regenerate the injured heart by blocking Cxcr4
function is caused by a defect in the CMs themselves.
DISCUSSION
Two cxcl12-cxcr4 systems in zebrafish
In this report, we identified cxcl12a-cxcr4b-dependent CM migration
as an essential mechanism for heart regeneration in zebrafish. The
DEVELOPMENT
4138 RESEARCH ARTICLE
Fig. 7. Cxcr4 function is required for CM migration during heart
regeneration. (A)Experimental strategy to assay CM migration during
heart regeneration. After localized photoconversion of the cmlc2aKaede heart, hearts are exposed by enlarging the pericardiac window,
and the ventricular apex is amputated. At the desired time, labeled
CMs are examined by imaging analysis. (B-O)Green signal represents
non-irradiated Kaede and newly synthesized Kaede after irradiation and
red signal represents photoconverted Kaede. C, E, I, M, G, K and O
show higher magnification images of boxed area in B, D, H, L, F, J and
N, respectively. (B,C)Immediately after photoconversion and
amputation, green signal was lost at the irradiated site (B, arrowheads),
where red signal was detected (C, arrows). Ventricular amputation was
performed in the non-irradiated area. (D,E)At 7 dpa, red signals
(photoconverted CMs, white arrowheads in E) and green signals (nonphotoconverted CMs, open arrowheads in E) were detected in the
injury site. (H-M)At 10 dpa (H,I) and 14 dpa (L,M), red signals and
green signals were also detected in the injury site. (F-O)CMs were not
detected in the injury site of the CXCR4-antagonist-treated heart. All
photoconverted CMs stayed outside the injury site (yellow arrows in
G,K,O) by blocking Cxcr4 function. (P)Quantitation of the migrated
CMs 7, 10 and 14 days after photoconversion and amputation. The
vertical axis represents the percentage of photoconverted CMs in the
injury site compared with the number of entire photoconverted CMs. A
section at the center of the injury site was examined from each heart.
Dotted lines indicate the amputation planes. The yellow arrowheads in
B, D and F point to the irradiated areas. The yellow arrows in C, E, G, I,
M, K and O point to red signals outside the injury site. The open
arrowheads and white arrowheads point to non-photoconverted CMs
and photoconverted CMs, respectively, in the regenerating area. The
asterisks in D, J and N indicate the valves between the atrium and
ventricle. Scale bars: 50m.
CXCL12-CXCR4 system is a major chemokine-receptor system that
regulates directed migration of a variety of cells (Raz and
Mahabaleshwar, 2009). Zebrafish have two cxcl12 genes and two
cxcr4 genes as a result of the teleost genome duplication during
RESEARCH ARTICLE 4139
Fig. 8. Normal neo-vascularization and activation of FGF
signaling after blocking Cxcr4 function. (A-D)fli1-EGFP (green) and
cmlc2a-mCherry (magenta) signal in the control (A,C) and CXCR4antagonist-treated (B,D) heart at 7 dpa (A,B) and 14 dpa (C,D).
Arrowheads point to neo-vascularization in the regenerating area,
visualized by the fli1-EGFP signal. (E)Quantitation of neovascularization. The fli-EGFP positive area compared with the
regenerating area was measured and by ImageJ software. A section at
the center of the injury site of each heart was examined (n3). (F-I)FGF
signaling status, visualized by phospho ERK1/2 (pERK1/2)
immunoreactivity (F,G) and expression of mkp3/dusp6 (H,I) at 14 dpa in
control (F,H) and CXCR4 antagonist-treated (G,I) hearts. The
arrowheads point to the pERK1/2 (F,G) and mkp3/dusp6 (H,I) signals.
The dotted lines indicate the amputation planes. Scale bars: 50m.
evolution (Amores et al., 1998). This gene duplication appears to
contribute to functional segregation of the cxcl12-cxcr4 system. For
instance, during embryonic development, the cxcl12b-cxcr4a system
functions for blood vessel development, endothelial cell migration
(Bussmann et al., 2011; Siekmann et al., 2009), and endoderm
migration during gastrulation (Mizoguchi et al., 2008; Nair and
Schilling, 2008). The cxcl12a-cxcr4b system is known to function
for primordial germ cell migration (Knaut et al., 2003; Raz, 2003),
sensory ganglia assembly (Knaut et al., 2005) and lateral line
migration (Haas and Gilmour, 2006), and is also expressed in
regenerating fins (Bouzaffour et al., 2009). This functional
segregation might have contributed to the viability of the cxcr4b
mutant line, because mouse mutants that lack either Cxcl12 or Cxcr4
die before birth owing to a ventricular septal defect, defective
formation of the large vessels in the gastrointestinal tract and
impaired hematopoietic development (Ma et al., 1998; Tachibana et
al., 1998; Zou et al., 1998). Our data demonstrated that the cxcl12acxcr4b system functions to regulate CM migration, which is essential
for heart regeneration, and that the cxcl12b-cxcr4a system might be
involved in the regeneration of other organs.
DEVELOPMENT
Cardiomyocyte migration
4140 RESEARCH ARTICLE
Cxcr4 function is necessary in CMs during heart
regeneration in zebrafish
Our analysis showed that Cxcr4 functions for directed migration of
CMs toward the injury site in zebrafish. This is in contrast to
mammalian myocardial infarction models (Takahashi, 2010), in
which Cxcr4 functions in CMs (Hu et al., 2007) and bone-marrowderived mesenchymal stromal cells (Honczarenko et al., 2006). In
mammals, Cxcr4 in CMs is shown to act for enhanced cell survival
and reduction of infarction size (Hu et al., 2007). The Cxcl12-Cxcr4
system in the mesenchymal stromal cells functions for
cardioprotection (Saxena et al., 2008), and a fraction of bonemarrow-derived cells can differentiate into CMs (Wojakowski et al.,
2010). In amputated zebrafish hearts, CM proliferation and survival
were not affected by blocking Cxcr4 function (Fig. 5; supplementary
material Fig. S7). Moreover, it is yet to be determined whether
mesenchymal stromal cells in zebrafish (Lund et al., 2012) can
contribute to CMs. Nonetheless, our data highlight the requirement
of Cxcr4 function in CMs during heart regeneration in zebrafish.
The Cxcl12-Cxcr4 system and neo-vascularization
Our analysis showed a lack of Cxcr4 signal in endothelial cells in
regenerating hearts (Fig. 2). This contrasts to mammalian
myocardial infarction models (Takahashi, 2010), according to
which Cxcr4 is expressed in cell types involved in neovascularization, such as bone-marrow-derived mesenchymal
stromal cells (Honczarenko et al., 2006; Yamaguchi et al., 2003)
and endothelial progenitor cells (Yamaguchi et al., 2003). Although
zebrafish stromal cells can exhibit endothelial-like properties (Lund
et al., 2012), neo-vascularization seems to be unaffected in
CXCR4-antagonist treated fish (Fig. 8). Studies have demonstrated
that neo-vascularization in regenerating zebrafish hearts involves
the contribution of epicardial cells (Kim et al., 2010; Lepilina et al.,
2006), which are unlikely to be affected by CXCR4 antagonist
treatment (supplementary material Fig. S9). The present study does
not rule out the possibility that other unidentified cell types migrate
to the injured heart in a Cxcr4-dependent manner and contribute to
heart regeneration in zebrafish. However, data obtained in our
analyses suggest that neo-vascularization occurs independently
from Cxcr4 function during heart regeneration in zebrafish (Fig. 9).
Kaede-photoconversion system and cell tracing
Cell lineage analysis is an important issue to understand complex
processes of regeneration, in which multiple cell types are involved
(Tanaka and Reddien, 2011). A genetic recombination approach
Development 139 (22)
using CreER transgenic lines is a powerful method, especially for
long-term lineage analysis (Jopling et al., 2010; Kikuchi et al.,
2010). The advent of photoconvertible fluorescent proteins, such
as Kaede, has led to the development of an effective approach for
tracing migration of specific cell types in vivo (Ando et al., 2002).
Our data show that CMs labeled by Kaede photoconversion
migrate toward the injury site during heart regeneration (Fig. 7).
Recent studies also showed neutrophil mobilization in zebrafish
larvae (Deng et al., 2011) and developmental timing assays during
zebrafish heart development (de Pater et al., 2009) with similar
approaches. Thus, localized photoconversion in combination with
the use of cell-type-specific promoters/enhancers would be a
valuable approach for cell migration and lineage analysis in a
variety of biological processes.
A role for CM migration during heart
regeneration
The present study demonstrates that CM migration is an essential
mechanism for heart regeneration, in addition to CM proliferation
and vasculature formation. Considering that clearing fibrin scarring
would also be important for heart regeneration, remodeling of the
extracellular matrix should also be coupled to these processes. Such
a process might involve unidentified molecular systems, and is to
be studied in the future. Our data also indicate that the proliferation
of CMs and neo-vascularization are regulated independently from
migration (Fig. 9). As both proliferation of CMs (Jopling et al.,
2010; Kikuchi et al., 2010) and their migration (this study) are
necessary, these two events need to be coordinated for the
regeneration of the injured heart. Previous studies show that genes
involved both in cell cycle regulation and cell movement are
upregulated in regenerating zebrafish hearts (Lien et al., 2006;
Sleep et al., 2010), which supports the conclusions of our research.
Given that the neonatal mammalian heart also possesses the ability
to regenerate after resectioning through the proliferation of preexisting CMs (Porrello et al., 2011), the correct coordination of
migration and proliferation may prove to be crucial for heart
regeneration not only in zebrafish but also in mammalian species.
Acknowledgements
We thank the zebrafish core facility at the University of Minnesota for general
help with the breeding and maintenance of zebrafish lines. We thank Drs
Michael O’Connor, Yasushi Nakagawa, Angel Raya, Erez Raz and Koichi
Kawakami for sharing equipment or materials. We thank Drs Jonathan Slack
and Naoko Koyano for critical reading, and the Developmental Studies
Hybridoma Bank developed under the auspices of the NICHD and maintained
by The University of Iowa.
Funding
I.O was supported by the Japan Society for the Promotion of Science, Grant-inAid for Scientific Research (C) [22570197], by the Mochida Memorial
Foundation for Medical and Pharmaceutical Research and by the Kowa Life
Science Foundation. Research in Y.K.’s lab was supported by the Minnesota
Medical Foundation [4099-9216-12].
Supplementary material
Supplementary material available online at
http://dev.biologists.org/lookup/suppl/doi:10.1242/dev.079756/-/DC1
Fig. 9. A model of heart regeneration in zebrafish. Amputation
induces cxcl12a-cxcr4b-dependent directed migration of CMs into the
injury site. CM proliferation and neo-vascularization are regulated
independently from CM migration. Coordinated progression of these
processes regenerates the injured heart. The orange circles represent
proliferating CMs.
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