ARF1 recruits RAC1 to leading edge in neutrophil chemotaxis
Mazaki et al. Cell Communication and Signaling
ARF1 recruits RAC1 to leading edge in neutrophil chemotaxis
Yuichi Mazaki 0
Yasuhito Onodera 1
Tsunehito Higashi 0
Takahiro Horinouchi 0
Tsukasa Oikawa 1
Hisataka Sabe 1
0 Department of Cellular Pharmacology, Graduate School of Medicine, Hokkaido University , Sapporo , Japan
1 Department of Molecular Biology, Graduate School of Medicine, Hokkaido University , Sapporo , Japan
Background: The small GTPase ARF1 mediates membrane trafficking mostly from the Golgi, and is essential for the G protein-coupled receptor (GPCR)-mediated chemotaxis of neutrophils. In this process, ARF1 is activated by the guanine nucleotide exchanger GBF1, and is inactivated by the GTPase-activating protein GIT2. Neutrophils generate the Gβγ-PAK1-αPIX-GIT2 linear complex during GPCR-induced chemotaxis, in which αPIX activates RAC1/CDC42, which then employs PAK1. However, it has remained unclear as to why GIT2 is included in this complex. Results: We investigated the association between ARF1 and RAC1/CDC42 during the fMLP-stimulated chemotaxis of HL60 cells. We found that the silencing of GBF1 significantly impaired the recruitment of RAC1 to the leading edges, but not PAK1, αPIX, RAC2, or CDC42. A significant population of RAC1 colocalized with ARF1 at the leading edges in stimulated cells, whereas fMLP activated both ARF1 and ARF5. Consistently, the silencing of ARF1, but not ARF5, impaired the recruitment of RAC1, whereas the silencing of RAC1 did not affect the recruitment of ARF1 to the leading edges. Conclusions: Our results indicated that the activation of ARF1 triggers the plasma membrane recruitment of RAC1 in GPCR-mediated chemotaxis, which is essential for cortical actin remodeling. Thus, membrane remodeling at the leading edges appears to precede actin remodeling in chemotaxis. Together with the fact that GIT2, which inactivates ARF1, is an integral component of the machinery activating RAC1, we proposed a model in which the ARF1-RAC1 linkage enables the regulation of ARF1 by repetitive on/off cycles during GPCR-mediated neutrophil chemotaxis.
Chemotaxis; ARF1; GBF1; RAC1
Neutrophils are rapidly polarized upon the detection of
a chemoattractant gradient, and start to migrate toward
the chemoattractant source. Such directional cell
migration requires a complex but well organized series of
intracellular events, such as cytoskeleton remodeling,
and membrane trafficking and remodeling. Most
chemoattractants, including N-formyl-Met-Leu-Phe (fMLP),
bind to their cognate G protein-coupled receptors
(GPCRs), and this binding then releases the Gα subunit
and the Gβγ heterodimer from heterotrimeric G
proteins to transmit the downstream signals [
]. αPIX is a
Dbl-family guanine nucleotide exchange factor (GEF) for
RAC1 and CDC42 , whereas p21-activating protein 1
(PAK1) is a downstream effector of activated RAC1 and
]. In neutrophil chemotaxis, Gβγ binds to
PAK1, which then binds to αPIX, thus forming the linear
complex of Gβγ-PAK1-αPIX, which regulates the
activities of the RHO-family GTPases, to remodel the
actinbased cytoskeletal structure upon GPCR signaling [
During GPCR-induced neutrophil chemotaxis, RAC1
primarily controls directional sensing, in which
RAC1deficient neutrophils frequently generate multi-head
leading edges during chemotaxis [
CDC42deficient neutrophils also generates similar multi-head
leading edges [
]. On the other hand, RAC2 appears to
be crucial for actin polymerization at the leading edges,
as RAC2-deficient neutrophils showed significant defects
in actin polymerization upon GPCR stimulation, and
thereby a loss of chemokinesis [
Membrane remodeling is another essential part of
neutrophil chemotaxis. ARF-family GTPases are
primarily engaged in membrane trafficking and remodeling,
and are hence crucial to higher order cellular functions,
(See figure on previous page.)
Fig. 1 Independence of GBF1 in the translocation of PAK1, αPIX, and Gβγ to the leading edges. (a) Expression pattern of proteins in cells treated
with GBF1 siRNAs . Cells transfected with siRNA against GBF1 or an irrelevant RNA duplex (Irr) were analyzed for expression of the indicated
proteins by immunoblotting of the lysates (10 μg each). Data are representative of three independent experiments. (b-g) Subcellular localization
of αPIX, PAK1, and Gβ. Differentiated HL-60 cells, transfected with GBF1 siRNA or Irr, were incubated with or without fMLP for 15 min, and
subjected to anti-αPIX (b), anti-PAK1 (d), or anti-Gβ immunostaining (f), and percentages of αPIX molecules (c), PAK1 molecules (e), or Gβ
molecules (g) translocated to the leading edges in fMLP-stimulated cells were calculated. F-actin was visualized by Texas Red-phalloidin. Data are
representative images of three independent experiments (b, d, and f), and >25 cells were analyzed in three independent experiments (c, e, and
g). Error bars, SEM (c, e, and g). ** p < 0.01 and * p < 0.05 compared with the Irr control. Bars, 10 μm (b, d, and f)
including cell motility [
]. ARF1 is primarily involved
in membrane and vesicle trafficking from the Golgi [
]. We previously showed that GIT2, which is a
GTPase-activating protein (GAP) for ARF1, binds to
αPIX to form a linear complex of Gβγ-PAK1-αPIX-GIT2
. This complex, as well as GIT2 on its own, was
crucial for the suppressive control of ARF1 activity during
GPCR signaling. Interestingly, GIT2 was moreover
found to be crucially involved in the efficient
recruitment and activation of RAC1 upon GPCR stimulation,
whereas CDC42 and RAC2 were almost unaffected by
the GIT2 deficiency [
]. As a result, GIT2-deficient
neutrophils lose their directional persistency in
GPCRmediated chemotaxis, whereas the rates of
actincytoskeletal polymerization and cell migration are almost
unaffected. Furthermore, the suppressive control of
ARF1 by GIT2 is important for the proper production of
superoxide, both in time and in direction, during
GPCRmediated neutrophil chemotaxis [
Processes activating ARF1 appear to be important for
several aspects of neutrophil chemotaxis. We identified
that among the different ARFGEFs, GBF1 is most crucial
for the activation of ARF1 in neutrophils upon GPCR
stimulation, in which GBF1 first translocates from the
Golgi to the leading edges, and then recruits ARF1 and
GIT2 to the leading edges [
]. In this process, the
expression of a dominant-active form of ARF1, namely
ARF1(Q71L), was sufficient to recruit GIT2. GBF1
silencing impaired the directional migration, whereas cell
migration rates were not notably affected; and moreover,
this silencing, as well as the expression of the
dominantnegative form of ARF1, ARF1(T31 N), frequently
generated multi-head leading edges during chemotaxis, similar
to those observed previously upon the deficiency of
RAC1 or CDC42 [
]. Thus, a close association appears
to exist between ARF1 and these RHO-family GTPases
in GPCR-mediated neutrophil chemotaxis, with regard
to their plasma membrane recruitment and activation.
Furthermore, GBF1 silencing was found to affect the
proper production of superoxide upon GPCR
stimulation, which might be a reflection of the fact that GBF1 is
required to recruit GIT2 to the leading edges [
ARF-family GTPases may function through their
cycles of activation and inactivation. For example,
expression of either the GTP hydrolysis-deficient mutant
or the GDP-bound mutant of ARF1 both blocked the
functions of ARF1 associated with ER-Golgi transport
]. However, the molecular mechanisms by which the
activation processes of the ARF-GTPases are coupled
with the inactivation processes remain unclear. We show
here that ARF1 activation recruits RAC1 to the leading
edges of GPCR-stimulated neutrophils, and propose that
this link generates a system in which ARF1 activation is
automatically coupled with its inactivation process at the
leading edges during GPCR-stimulated chemotaxis of
Results and Discussion
Recruitment of Gβγ, αPIX, and PAK1 to leading edges
occur independent of the GBF1
Silencing of GBF1 in HL-60 cells frequently generated
multi-head leading edges during fMLP-induced
chemotaxis, similar to those observed upon the inhibition of
RAC1 or CDC42 [
]. We have shown that GBF1 small
interfering RNA (siRNA) treatment causes loss of the
polarized accumulation of GIT2 at the leading edges of
fMLP-stimulated HL-60 cells, in which a greater than
50% decrease in the accumulation of GIT2 at actin-rich
leading edges was observed compared with cells treated
with a control irrelevant siRNA [
]. GIT2 forms a
complex with αPIX and PAK1, which are an activator and an
effector of RAC1, respectively. αPIX and PAK1
accumulate at actin-rich leading edges upon fMLP stimulation,
whereas these proteins mostly localize around the cell
periphery in unstimulated neutrophils [
] (also see
Fig. 1b and d). We hypothesized that the lack of GIT2
recruitment upon GBF1 silencing may impair the
recruitment of αPIX and PAK1, thus causing the
dysfunction of RAC1 at the leading edges. We then suppressed
the expression of GBF1 protein by siRNA method in
differentiated HL-60 cells. We found that two siRNA
sequences of GBF1 block expression of GBF1 protein
without notable suppression of others protein
expression in differentiated HL-60 cells (Fig. 1a). However,
unlike in the case of GIT2, GBF1 silencing decreased
the accumulation of αPIX and PAK1 at the leading
edges only by approximately 10% in fMLP-stimulated
HL-60 cells (Fig. 1b-e).
(See figure on previous page.)
Fig. 2 Requirement of GBF1 in RAC1 activity and its translocation to the leading edges. (a-f) Subcellular localization of CDC42, RAC1, or RAC2.
Differentiated HL-60 cells, transfected with GBF1 siRNA or an irrelevant RNA duplex (Irr), were incubated with or without fMLP for 15 min, and
subjected to anti-CDC42 (a), anti-RAC1 (c), or anti-RAC2 immunostaining (e), and percentages of CDC42 molecules (b), RAC1 molecules (d), and
RAC2 molecules (f) translocated to the leading edge in fMLP-stimulated cells were calculated. F-actin was visualized by Texas Red-phalloidin. Data
are representative images of three independent experiments (a, c, and e), and >25 cells were analyzed in three independent experiments (b, d,
and f). (g and h) Activities of CDC42, RAC1, and RAC2. Activities of CDC42, RAC1, and RAC2 were measured by GST-PBD pulldown coupled with
the indicated antibodies. Each lower panel represents immunoblots of total cell lysates (5 μg) by the indicated antibodies. Data are representative
of three independent experiments (g), and were analyzed in three independent experiments (h). In h, values for Irr control at 0 s are considered
1. Error bars, SEM (b, d, f and h). ** represents a statistical difference from the Irr control (p < 0.01). Bars, 10 μm (a, c, and e)
Gβγ forms a complex with GIT2, via αPIX and PAK1;
GIT2 deficiency caused a substantial loss of the
polarized accumulation of Gβγ at the leading edges of
GPCRstimulated neutrophils (> 50% decrease, Mazaki et al.,
2006). On the other hand, a substantial fraction of Gβγ
appeared to localize to the cytosol, rather than to the
cell periphery, in the unstimulated neutrophils [
see Fig. 1f ). No significant reduction was observed by
GBF1 silencing in the recruitment of Gβγ leading edges
upon fMLP stimulation (Fig. 1f and g). Collectively, it is
likely that although GBF1 is crucial for the recruitment
of GIT2 to leading edges upon GPCR signaling, the
recruitment of Gβγ, αPIX, and PAK1 to leading edges is
substantially independent of the GBF1-GIT2 axis,
despite the fact that these three proteins form a complex
with GIT2 in GPCR signaling.
GBF1 is required for recruitment and activation of RAC1
RAC1 and CDC42 are also recruited to the leading
edges upon GPCR stimulation of neutrophils [
whereas these proteins mostly localized around the cell
periphery in unstimulated cells (Fig. 2a and c). We next
examined the effects of GBF1 silencing on these
proteins, and found that the silencing of GBF1 significantly
impaired the recruitment of RAC1 to the leading edges
(decreased by 30%–40%), but not CDC42 (Fig. 2a-d).
GBF1 silencing caused no significant reduction in RAC2
accumulation at the leading edges (Fig. 2e and f ).
Moreover, GBF1 silencing significantly suppressed the
fMLPinduced activation of RAC1 in HL-60 cells, as measured
30 s after the stimulation, whereas this silencing did not
notably affect the activation of CDC42 and RAC2 (Fig.
2g and h). These results indicated that GBF1 is linked to
RAC1, rather than RAC2 or CDC42, in the GPCR
signaling of neutrophils.
ARF1 is required for recruitment and activation of RAC1
We then addressed the mechanism by which GBF1 is
involved in the recruitment of RAC1 to the leading edges.
ARF1 may not be the sole target of GBF1 [
fact, in addition to ARF1, fMLP stimulation activated
ARF5 in HL-60 cells (these cells express ARF1, 3, 4, and
5) (Fig. 3a and b). We then investigated the possible
colocalization of ARF1 and ARF5 with RAC1 upon
fMLP stimulation. ARF1 and RAC1 each showed a
punctate distribution at the leading edges of stimulated
HL-60 cells, in which a significant fraction of ARF1 and
RAC1 are colocalized (Fig. 3c and d). ARF5 also showed
a punctate distribution at the leading edges, but its
colocalization with RAC1 appeared to be very limited
compared with that of ARF1 (Fig. 3c and d). We then found
that silencing of ARF1 significantly inhibited the
recruitment of RAC1 to the leading edges upon fMLP
stimulation of HL-60 cells (~40% inhibition), whereas the
silencing of ARF5 did not affect RAC1 recruitment at all
(Fig. 3f and g). Thus, ARF1, but not ARF5, appeared to
be crucial for the recruitment of RAC1, whereas ARF1
and ARF5 are both activated under this condition.
Moreover, ARF1 silencing significantly suppressed the
fMLP-induced activation of RAC1 in HL-60 cells, as
measured 30 s after the stimulation (Fig. 3h and i). We
also analyzed whether the silencing of RAC1 affects the
recruitment of ARF1 to the leading edges upon fMLP
stimulation, and found that RAC1 was not at all
required for the recruitment of ARF1 (Fig. 3k and l).
Taken together, it is conceivable that the activation of
ARF1 by GBF1 plays an important role in the
recruitment of RAC1 to the leading edges and the activation of
RAC1 upon the GPCR signaling of neutrophils.
The fact that GIT2, which is a GAP for ARF1, is
an integral component of the Gβγ-PAK1-αPIX
] has been enigmatic, as this complex
apparently forms primarily to activate and assist in the
functioning of RAC1 and/or CDC42. In the present
study, we found that activation of ARF1 by GBF1 at
the leading edges of cells recruits RAC1 to the same
leading edges; we hence propose a model in which
ARF1 activity is regulated by repetitive on/off cycles
during GPCR-mediated neutrophil chemotaxis
(Fig 4). Our model explains that the inclusion of
GIT2 as a member of the Gβγ-PAK1-αPIX complex
provides a system by which ARF1 activity can be
repetitively regulated between its activation and
inactivation cycles, concurrently with plasma
membrane protrusion and the formation of leading
edges. In other words, ARF1 on its own appears to
generate this system by recruiting RAC1, in order
to perform directional cell migration.
(See figure on previous page.)
Fig. 3 Requirement of ARF1 in RAC1 activity and its translocation to the leading edges. (a and b) Activity of ARFs in differentiated HL-60 cells after
fMLP stimulation. Activities of class I and II ARFs were measured by GST-GGA3 pulldown coupled with the indicated antibodies. Each lower panel
represents immunoblots of the total cell lysates (5 μg) by the indicated antibodies. Data are representative of three independent experiments (a),
and were analyzed in three independent experiments (b). In b, values of each GTP-ARF at 0 s are considered 1. ** p < 0.01 and * p < 0.05
compared with each GTP-ARF at 0 s. (c and d) Subcellular localization of RAC1, ARF1, and ARF5 after fMLP stimulation. Differentiated HL-60 cells were
incubated with fMLP for 5 min, and subjected to immunostaining analysis, using high-resolution SIM. Specificities of the anti-ARF1 antibody and
the anti-RAC1 antibody were confirmed by ARF1 or RAC1 siRNA-treatment of cells (c) Bars, 2 μm. Pearson’s correlation coefficients of the
intracellular colocalization of these proteins, as indicated, were estimated from >10 cells (d). (e) Suppression of ARF1 or ARF5 by siRNAs in differentiated
HL-60 cells. Cells transfected with siRNA against ARF1, ARF5, or an irrelevant RNA duplex (Irr) were analyzed for the expression of the indicated
proteins by immunoblotting of the lysates (10 μg each). Data are representative of three independent experiments. (f and g) Subcellular
localization of RAC1. Differentiated HL-60 cells, transfected with siRNA against ARF1, ARF5, or Irr, were incubated with fMLP for 15 min, and
subjected to anti-RAC1 immunostaining (f), and percentages of RAC1 molecules translocated to the leading edges in fMLP-stimulated cells were
calculated (g). (h and i) Activities of RAC1. Activities of RAC1 were measured by GST-PBD pulldown coupled with the anti-RAC1 antibodies. Each
lower panel represents immunoblots of total cell lysates (5 μg) by the anti-RAC1 antibodies. Data are representative of three independent
experiments (h), and were analyzed in three independent experiments (i). In i, values for Irr control at 0 s are considered 1. (j) Suppression of
RAC1 by siRNAs in differentiated HL-60 cells. Cells transfected with siRNA against RAC1 or Irr were analyzed for expression of the indicated
proteins, by immunoblotting of the lysates (10 μg each). Data are representative of three independent experiments. (k and l) Subcellular
localization of ARF1. Differentiated HL-60 cells, transfected with RAC1 siRNA or Irr, were incubated with or without fMLP, as indicated, and
subjected to anti-ARF1 immunostaining (k), and percentages of ARF1 molecules translocated to the leading edges in fMLP-stimulated cells were
calculated (l). F-actin was visualized by Texas Red-phalloidin. Data are representative images of three independent experiments (f, and k), and >25
cells were analyzed in three independent experiments (g and l). Error bars, SEM (b, d, g, i and l). ** represents a statistical difference from Irr
(p < 0.01) (g and i). Bars, 10 μm (f and k)
Our results demonstrated that ARF1 activation precedes
RAC1 activation and function in cell migration. This
notion is consistent with the concept previously proposed by
Donaldson, in which ARF-mediated membrane
remodeling is a prerequisite for actin-cytoskeletal remodeling,
which is mediated by RHO-family GTPases [
ARF1 might not always be inactivated when RAC1 is
activated. Wiskott-Aldrich syndrome protein
(WASP)family verprolin homologous protein (WAVE) regulatory
complex (WRC) is crucial for the dynamic regulation of
the structure of leading edges, by promoting membrane
ruffling and lamellipodia formation [
RAC1 is essential for WRC function [
]. It was
reported that the binding affinity of GTP-RAC1 to WRC
is relatively low, and that although GTP-ARF1 also binds
weakly to WRC, GTP-ARF1 assists the binding of RAC1
to WRC [
]. Consistently, cooperation of ARF1 and
RAC1 was shown to be necessary to induce
WAVEinduced actin polymerization [
]. Thus, our results may
have also illustrated a process in which these two small
GTPases are both activated and function cooperatively
with each other.
Then, an important question remains as to how the
timing of the inactivation of ARF1 by the
Gβγ-PAK1αPIX-GIT2 complex is regulated. αPIX can bind directly
to the plasma membrane via its pleckstrin homology
domain, and this binding recruits PAK1 to the plasma
]. As αPIX and PAK1, as well as GIT2,
are accumulated at leading edges upon GPCR
stimulation, it is likely that the Gβγ-PAK1-αPIX-GIT2 complex
is formed at the leading edges. On the other hand,
activation of ARF1 by GBF1 occurs independently of this
complex upon GPCR stimulation, as we have shown that
GBF1 is activated by a product of
phosphatidylinositol3-phosphate kinase γ (PI3Kγ) , which is activated via
its binding to Gβγ [
]. Thus, a more precise picture of
the inactivation process of ARF1 by the
Gβγ-PAK1αPIX-GIT2 complex will be required to understand the
nature of neutrophil chemotaxis. Mechanisms by which
ARF1 recruits RAC1 also await to be clarified.
Materials and methods
HL-60 cells were obtained from ATCC. HL-60 cells were
cultured in RPMI 1640 medium supplemented with 10%
fetal bovine serum (Gibco) and 2 mM L-glutamine. For
differentiation into neutrophil-like cells, cells were
cultured in the presence of 1.25% dimethyl sulfoxide for
6 days, as described previously [
Antibodies and chemicals
Antibodies were purchased from the following
commercial sources: mouse monoclonal antibody against ARF1
and ARF5 (Abcam), RAC1 (Millipore), CDC42 (Santa
Cruz Biotechnology), αPIX (Abnova), actin (Sigma);
rabbit polyclonal antibodies against ARF3, ARF4, and
RAC1 (Abcam), RAC2 and Gβ (Millipore), and PAK1
(Santa Cruz Biotechnology). Donkey antibody against
mouse and rabbit IgG, conjugated with horseradish
peroxidase, were from Jackson ImmunoResearch
Laboratories. Goat antibodies against mouse and rabbit IgGs,
conjugated with Alexa Fluor 488 or Alexa Fluor 555,
and phalloidins, conjugated with Texas Red, were from
Invitrogen. All other chemical reagents were purchased
from Sigma and Nacalai, unless otherwise stated.
Transfections were performed as described previously [
For the transfection of siRNA, 3 μg of siRNAs each specific
to GBF1, ARF1, ARF5, and RAC1, or an irrelevant RNA
duplex (siCONTROL, RISC-free siRNA1; Dharmacon) were
used. GBF1 siRNA targeting sequences were as described
]. ARF1 and ARF5 siRNA targeting sequences
were 5′-TGACAGAGAGCGTGTGAAC-3′ (ARF1 sequence
1), 5′-ACCGUGGAGUACAAGAACA-3′ (ARF1 sequence
2), 5′- TCTGCTGATGAACTCCAGA-3′ (ARF5 sequence
1) and 5′- CCATAGGCTTCAATGTAGA-3′ (ARF5
sequence 2), as described previously [
]. RAC1 siRNA
targeting sequences were 5′- AGACGGAGCTGTAGGTAAA-3′
(RAC1 sequence 1) and
5′-TAAGGAGATTGGTGCTGTA3′ (RAC1 sequence 2), as described previously [
Immunofluorescence microscopy was performed as
described previously [
]. Briefly, differentiated HL-60 cells
were attached to coverslips in Hank’s balanced salt
solution (HBSS) containing 20 mM
4-(2-hydroxyethyl)-1piperazineethanesulfonic acid (pH 7.2) and 0.1% bovine
serum albumin. Coverslips were then placed on Dunn
chambers (Hawksley), and incubated for the indicated
times at 37 °C. Acquisition of confocal images using a
laser-scanning microscope (FV500; Olympus) was
performed as previously described [
]. Each experiment was
performed three times, in each of which more than 50
cells were analyzed, and representative images are shown
in each figure. High-resolution structured illumination
(SIM) microscopy analysis was performed to analyze the
intracellular colocalization of RAC1 with ARF1 or with
ARF5 using an N-SIM microscope (Nikon) and
NISelements software (Nikon), as described previously [
Small GTPase activities
For measurement of small GTPase activities, 1 × 106
cells were washed, and preincubated in HBSS for 5 min
at 37 °C, and then stimulated with 100 nM fMLP or left
untreated for the indicated times in the same solution at
37 °C. Cells were then solubilized, and GTP-bound class
I and II ARFs were pulled-down using 50 μg of
glutathione S-transferase (GST)-GGA31–226 [
CDC42, RAC1, and RAC2 were pulled-down using
]. Amounts of these GTPases in total cell
lysates were simultaneously determined by
immunoblotting using their antibodies. Small GTPase activities were
measured by a densitometer (GT-X770 scanner; Epson)
using Image version 1.50i software (National Institutes
of Health, Bethesda, MD).
For all experiments, differences between groups were
calculated by Tukey-Kramer test.
fMLP: N-formyl-Met-Leu-Phe peptide; GAP: GTPase-activating protein;
GEF: Guanine nucleotide exchanging factor; GPCR: G protein-coupled
receptor; GST: Glutathione S-transferase; PAK1: p21-activating protein kinase 1;
SIM: Structured illumination microscopy; siRNA: small interfering RNA;
WRC: WAVE regulatory complex
We thank A. Hirano and E. Hayashi for their assistance, and H. A. Popiel for
critical reading of the manuscript. We also thank Nikon Instech Co. for their
help with the SIM analysis.
This work was supported by grants-in-aid from the Ministry of Education,
Science, Sports and Culture of Japan, grants from Novartis Foundation for the
Promotion of Science. Y. M. was partially supported by Special Coordination
Funds for Promoting Science and Technology from the Japan Science and
Availability of data and materials
All data used in this study are available from the corresponding author on
YM and HS contributed to the design of the study. YM, YO, T. Higashi, T.
Horinouchi and TO performed the experiments and analyzed the data. YM
and HS wrote the manuscript. All authors read and approved the final
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
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