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First published online August 24, 2007
doi: 10.1242/10.1242/dev.02883
1 Department of Molecular Physiology and Biophysics, Baylor College of Medicine,
Houston, TX 77030, USA.
2 Department of Chemical Engineering, California Institute of Technology,
Pasadena, CA 91125, USA.
3 Department of Medicine, School of Medicine, University of California-San
Diego, 9500 Gilman Drive, La Jolla, CA 92093-0641, USA.
4 Biological Imaging Center, Department of Biology, California Institute of
Technology, Pasadena, CA 91125, USA.
Author for correspondence (e-mail:
mdickins{at}bcm.tmc.edu)
Accepted 5 July 2007
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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Key words: Blood flow, Viscosity, Shear stress, Cardiovascular, Angiogenesis, eNOS (Nos3), Mlc2a (Myl7), Mouse
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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; Ji et al.,
2003; Tam and Schoenwolf,
1999). Concurrent with heart formation, blood and vessels first
form in the extraembryonic yolk sac. By 8.5 days post-coitum (dpc), a
capillary plexus consisting of a polygonal network of endothelial-lined
channels is evident in the yolk sac and these vessels ultimately connect to
the embryo, completing a circulatory loop between embryonic and
extra-embryonic tissues. Once the heart begins to beat, primitive
erythroblasts originating in the yolk sac can be found in the embryo by 4- to
6-somites (Ji et al., 2003;
McGrath et al., 2003). Between
8.5 and 9.5 dpc, primitive vessels of the yolk sac undergo remodeling in the
presence of flow to form a branched, hierarchical network of large and
small-caliber vessels surrounded by smooth muscle cells (for a review, see
Armulik et al., 2005).
The normal development of the cardiovascular system depends on a large
number of genes, suggesting complex signaling pathways
(Argraves and Drake, 2005;
Solloway and Harvey, 2003;
Trinh and Stainier, 2004). In
fact, just one aspect of development - vascular remodeling in the murine yolk
sac - requires over 60 known genes (our unpublished observations), including
members of the Tgf-ß, Notch, Vegf, hedgehog and retinoic acid signaling
pathways. Although many genes required for proper vascular remodeling are
expressed in endothelial cells and are thought to play a role in these cells
directly, mutations in many genes required solely for cardiac function also
cause defects in vascular remodeling. For instance, null mutations in
Ncx1 (also known as Slc8a1 - Mouse Genome Informatics)
(Koushik et al., 2001;
Wakimoto et al., 2000), myosin
light chain 2a (Mlc2a; also known as Myl7 - Mouse Genome
Informatics) (Huang et al.,
2003), Nkx2.5 (Tanaka
et al., 1999) and titin (May
et al., 2004) all exhibit failed yolk sac remodeling despite a
lack of expression of these genes in endothelial cells. Furthermore, vascular
abnormalities caused by the deletion of N-cadherin (cadherin 2) can be rescued
by cardiac-specific expression of either N- or E-cadherin (cadherin 1)
(Luo et al., 2001). To explain
these effects, it has been proposed that normal blood flow is necessary for
vascular remodeling. Numerous studies from avian embryos, dating back to well
over a century ago, support this hypothesis because the surgical manipulation
of blood flow can lead to a wide range of abnormalities in heart and vessel
development (Keller, 2001;
Kurz, 2000).
Whereas many studies show that blood flow is required for vessel and
cardiac remodeling, what continues to be debated is the reason why blood flow
is important (Conway et al.,
2003; Coultas et al.,
2005; Huang et al.,
2003; May et al.,
2004). For instance, it can be argued that blood flow is required
to deliver oxygen or nutrients to remodeling tissues. Hypoxic conditions could
damage endothelial cells directly or alter the levels of necessary growth
factors such as Vegf (also known as Vegfa - Mouse Genome Informatics), which
could impair remodeling (Conway et al.,
2003; May et al.,
2004). However, early mouse embryos (8.5 dpc), like early chick
(Cirotto and Arangi, 1989;
Pelster and Burggren, 1996;
Territo and Burggren, 1998),
frog (Territo and Burggren,
1998) and zebrafish embryos
(Pelster and Burggren, 1996),
can be cultured in the presence of carbon monoxide, which competes for oxygen
binding to hemoglobin, without effecting the initial stages of vasculogenesis
or vascular remodeling (E.A.V.J., S.E.F. and M.E.D., unpublished). Another
argument is that early blood flow functions to transport soluble nutrients,
such as cell-signaling molecules that are required for vascular remodeling
(Conway et al., 2003), although
there is no direct evidence to support such a mechanism.
A third hypothesis for the role of circulation is that it imparts force on
plexus endothelial cells and this mechanical force is necessary to activate
cell-signaling cascades necessary for vessel remodeling. Blood flow exerts two
types of force on vessels: shear stress, the frictional force tangential to
endothelial cells, and circumferential strain, a force perpendicular to the
direction of flow and which is related to intravascular pressure. Numerous
studies have shown that cultured endothelial cells can respond to these forces
by changing morphology, activating intracellular kinases and inducing gene
expression (for a review, see Li et al.,
2005) and shear stress levels in the mouse yolk sac are within the
range known to induce these responses in cultured endothelial cells
(Jones et al., 2004). Despite
the circumstantial evidence for flow-derived force in vascular remodeling, no
direct evidence exists to show that mechanical signals drive remodeling during
development or to exclude the possibility that the local oxygen supply or
soluble paracrine factors provided through circulation act as a trigger.
To better understand the role of circulation during early embryonic mouse
development, we have used whole-embryo culture and time-lapse confocal imaging
(Jones et al., 2002) to
visualize and quantify blood flow in early embryos. We first used these
methods to analyze dynamic events at the onset of circulation in normal mouse
embryos. Next, we characterized circulation deficiencies in
Mlc2a-null embryos to understand how secondary remodeling defects are
generated by impaired cardiac contractility. Finally, we tested whether
altering the mechanical properties of the early blood can phenocopy the
vascular remodeling defect seen in Mlc2a-/- embryos. These
studies reveal that erythroblast circulation begins after a prolonged period
of plasma flow, that both plasma and erythroblast flow are disrupted in
Mlc2a mutant embryos, and that the change in effective viscosity
caused by the entry of blood cells into circulation is required to induce
proper remodeling. Thus, using novel methods to analyze and manipulate blood
flow in early mouse embryos, we have answered a century-old question about the
role of mechanical force in vessel remodeling in vivo.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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-globin-GFP) mice or
Tg(-globin-GFP) mice
(Dyer et al., 2001;
Huang et al., 2003) were mated
and the presence of a vaginal plug was taken as 0.5 dpc. Embryos were
collected at 8.5 dpc and cultured for time-lapse microscopy
(Jones et al., 2002). To study
the initiation of flow in normal embryos, single images were taken every 6
minutes using a 20x Plan-Neofluar 0.5NA lens on a Zeiss LSM 5 PASCAL.
For tracking erythroblast motion in Mlc2a mutants, images were taken
at two frames/second using a 40x C-Apochromat 1.2NA objective.
Individual cells were tracked manually by connecting the center of the same
cell in subsequent images. Overlay of tracking data onto the images was
performed using Adobe Photoshop.
Plasma flow measurements
To determine if plasma could be transported from the embryo to the yolk
sac, 0- to 6-somite embryos were dissected as previously described
(Jones et al., 2002). A pulled
quartz needle (Sutter Instruments) was filled with 10x103
Mr fluorescein-dextran (Molecular Probes, D-1821) and a
picospritzer II (General Valve Corporation) was used to inject nanoliter
volumes of dye into the heart tube. Embryos were allowed to recover for up to
10 minutes at 37°C and were imaged with the Zeiss LSM PASCAL at 20x
magnification to determination whether plasma was transported to the yolk
sac.
For FRAP experiments, embryos at different stages were injected with
10x103 Mr fluorescein-dextran (50 mg/ml)
and transferred to Nunc chambers (No. 155380) with culture medium
(Jones et al., 2002). Embryos
recovered for 15 minutes to 1 hour at 37°C. The microscope stage (Zeiss
LSM 5 PASCAL or Zeiss LSM 510 META) was preheated to 37°C using a heater
box (Jones et al., 2002). For
reproducibility, measurements were always taken from the same region on the
arterial side of the yolk sac near the caudal end of the embryo. Since the
Mlc2a mutants cannot be unambiguously identified at the 8- to
9-somite stage, all embryos were measured blind and then genotyped after FRAP
experiments. Embryos with blood cells throughout the plexus were avoided so
that the movement of blood cells did not interfere with plasma measurements.
For acrylamide-treated embryos (see below), acrylamide was polymerized by the
addition of TEMED and embryos were allowed to recover at 37°C for 15-30
minutes prior to the injection of fluorescein-dextran (see Fig. S1 in the
supplementary material). Using a 20x Plan-Apochromat 0.75 NA lens, image
intensity was recorded within a region of interest (ROI) using 5% laser power.
Laser power was then increased to 100% and the ROI was scanned repeatedly to
bleach the fluorescence. Laser power was then reduced back to 5% to image the
recovery of the dye into the bleached field. Somites were counted following
the measurement.
Mean fluorescence with respect to time for the ROI was exported to a
spreadsheet. The recovery curve was fitted to the equation
(Soumpasis, 1983):
 |
where F(t) is the fluorescence intensity, F0
is the initial post-bleach fluorescence intensity, FF is
the final level of fluorescence recovery, I0 and
I1 are zeroth and first order Bessel's functions, and
is the characteristic diffusion time. The bleach area was divided by
characteristic diffusion time to give the measured diffusion. The baseline for
pure diffusion was obtained by injecting embryos with fluorescein-dextran as
described, stopping the heart by chilling the embryos at 4°C for 1 hour,
reheating the embryos to 37°C and performing FRAP.
Erythroblast immobilization
The blood islands of 5-somite stage Tg(-globin-GFP)
embryos were gently microinjected with a 30% (w/v) solution of
acrylamide:bis-acrylamide (37.5:1; 161-0158, BioRad) diluted 1:1 with
2xPBS; ammonium persulfate (161-0700, BioRad) was added to a final
concentration of 10 mM (see Fig. S1 in the supplementary material). The medium
was exchanged and the embryos recovered for approximately 15 minutes. To
catalyze polymerization of acrylamide, TEMED (161-0801, BioRad) was diluted
1:1 with 2xPBS and applied to the blood islands using a micropipette
with a low positive pressure. The tip of the micropipette was gently touched
to the surface of the blood islands at a number of points to initiate gel
formation. The optimal dilutions for both TEMED and acrylamide were determined
by dilution series (data not shown). The medium was subsequently changed and
the embryos placed in roller culture as previously described
(Tam, 1998). Three sets of
controls were performed: (1) uninjected embryos; (2) embryos injected only
with the acrylamide solution; and (3) embryos in which only TEMED was
delivered to the blood islands.
To assess embryo development, yolk sac growth and remodeling, embryos were imaged at the same magnification with a Zeiss Axiocam mounted on a Zeiss Lumar stereomicroscope equipped for fluorescence imaging. Texas Red-dextran (Molecular Probes, D-1828) was injected to better visualize remodeling. The image scale was calibrated to an etched glass standard and the yolk sac perimeter was traced using Zeiss Axiovision software. The area of the field within the traced image of the yolk sac was recorded as the size of the yolk sac. Turning was scored on a scale of 1 to 5 (1, not turned; 3, partially turned; 5, fully turned) and yolk sac vessel remodeling was scored on a scale of 1 to 5 (1, regular polygonal structure throughout yolk sac; 3, increased avascular space and lengthening of vascular segments but no obvious branching pattern; 5, hierarchical branching pattern present throughout the yolk sac with the appearance of large-caliber vessels). Embryos were not matched to their experimental group until after scoring. Statistics were performed using SPSS software (SPSS, Chicago, IL). Data were compared using one-way ANOVA with Bonferroni post-hoc analysis. Statistical significance was determined at a level of 5% alpha error between experimental groups and control for a single measure (P<0.05).
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Immunohistochemistry
Embryos were treated as described above to produce control, low-hematocrit
and low-hematocrit+hetastarch groups. After 24 hours in roller culture,
embryos were immediately fixed in 4% paraformaldehyde for 1 hour at 4°C
then transferred to methanol or PBS for storage. Yolk sacs were incubated
overnight at 4°C in one of the following primary antibodies: anti-PECAM-1
(Pinter et al., 2001),
anti-VE-cadherin (R&D Systems, Minneapolis, MN; 1:100), or anti-eNOS
(Santa Cruz Biotechnology, Santa Cruz, CA; 1:200). Alexa Fluor 594 or 568
secondary antibodies (Invitrogen, Carlsbad, CA) were used depending on the
primary species. Images of staining patterns were collected using a Zeiss 510
META confocal system and a 40x C-Apochromat 1.2NA objective. All images
for a single antibody were collected using the same laser power and detector
gain settings in order to visually compare the relative intensity levels.
Images shown are a maximum intensity projection of several optical slices.
Each image is a representative sample of the 5-11 embryos that were stained
from each group.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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;
McGrath et al., 2003). To
observe the onset of circulation in live embryos, we imaged the movement of
GFP-labeled erythroblasts using time-lapse confocal microscopy
(Jones et al., 2002).
Tg(-globin-eGFP) embryos (n=3) were placed in
culture at the 5- to 6-somite stage (Fig.
1 and see Movie 1 in the supplementary material). During the first
hour, the heart beat was evident but circulating erythroblasts were not
observed (Fig. 1A,H).
Occasionally, GFP+ erythroblasts were seen outside the blood islands, but only
as single, adherent, stationary cells (Fig.
1A, red arrow). It is not clear whether these cells differentiated
at these sites or were previously moving but had stopped. As erythroblasts
begin to enter the circulation, the volume percentage of erythroblasts in
vessels (hematocrit) is initially low and fluctuates from one frame to another
(Fig. 1B). Many blood cells
were seen to flow with a net forward motion, but some individual erythroblasts
became stationary and grouped together (see Movie 1 in the supplementary
material). Some groups of erythroblasts remained adherent to one another for
up to 3 hours (Fig. 1, arrows)
before dissociating and then rejoining the circulation. Thus, intermittent
erythroblast motion was detected as early as the 6- to 7-somite stage and
progressively more erythroblasts became recruited into the circulation over
the next few hours.
The onset of circulation is usually defined by the movement of
erythroblasts, but we were interested in determining whether significant
plasma flow exists prior to erythroblast circulation. In agreement with
previous findings, we detected a beating heart in embryos as early as the
3-somite stage (Navaratnam et al.,
1986). To test whether the heart could pump plasma from the embryo
to the yolk sac, we injected fluorescent dextran into the hearts of
early-somite stage embryos and examined whether the dextran could be observed
in the yolk sac plexus after a 10-minute recovery period. In 0- and 1-somite
stage embryos, injected dextran remained confined to the heart (n=4,
data not shown). Similarly, dextran did not distribute in most 2-somite stage
embryos (5 of 6 embryos, Fig.
2A,B). However, dextran was found consistently throughout the
capillary plexus including the blood islands in 3-somite and older embryos (20
of 20 embryos; Fig. 2C,D).
Higher magnification views of 3- and 6-somite
(Fig. 2E,F) yolk sacs are shown
to illustrate the distribution of the dextran within the plexus vessels. These
results show that a continuous vessel network between the capillary plexus is
established at least by the time the heart begins to beat.
The rapid distribution of dextran throughout the plexus suggested that there is significant plasma flow produced by the first heart beats. To confirm this, we measured plasma flow in embryos prior to the entry of erythroblasts using fluorescence recovery after photobleaching (FRAP). FRAP is used to measure the mobility of fluorescently tagged molecules by bleaching these molecules and recording how fast fluorescence intensity returns to the bleached area. The diffusion rate of 10x103 Mr fluorescein-dextran injected into the plexus was found to be 51±7.6 µm2/second (n=8) in the absence of heart function (see Materials and methods). Fluorescence recovery was significantly faster than diffusion in embryos with normal circulation and we have used the term perfusion coefficient to describe these measurements of flow. At the 3- to 4-somite stage, perfusion coefficients ranged between 100 and 500 µm2/second, indicating the presence of slow, but significant, flow (Fig. 3). The variability of plasma flow rates is not surprising given that the plexus is an extensive network of highly branched vessels with varying resistance to flow. Perfusion values increased in later-stage embryos (5- to 6-somite stage) and plasma flow in some regions was too fast to measure (Fig. 3, indicated by asterisk with arrow). Thus, the heart is a functional pump as soon as the myocardium begins to contract and there is a significant period of plasma flow that precedes the entry of erythroblasts into circulation.
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), making this mutant an excellent model to study the
relationship between circulation and remodeling. To visualize erythroblasts,
Mlc2a mutant mice were crossed with
Tg(-globin-eGFP) mice and fluorescence images of fixed
Mlc2a-/-; Tg(-globin-eGFP) embryos
confirmed that vascular remodeling was impaired
(Fig. 4).
Mlc2a-/-; Tg(-globin-eGFP) embryos
were indistinguishable from littermates at 8.5 dpc. However, 9.5 dpc
Mlc2a-/- yolk sac vessels retained the unremodeled
capillary plexus, whereas Mlc2a+/+ and
Mlc2a+/- vessels were remodeled. Although 8.5 dpc Mlc2a-/- embryos appear similar to littermates, we hypothesized that altered flow patterns at 8.5 dpc could be responsible for the lack of remodeling observed at 9.5 dpc. To determine whether circulation is delayed in Mlc2a-/- embryos, we compared the distribution of erythroblasts in Mlc2a mutant yolk sacs with heterozygote or wild-type embryos (Table 1). By the 10-somite stage, erythroblasts were routinely found throughout the plexus in wild-type and heterozygous embryos, but were not seen consistently throughout the whole yolk sac plexus until the 12- to 13-somite stage in the Mlc2a-/- embryos. Thus, erythroblast circulation appeared to be somewhat delayed in Mlc2a-/- embryos.
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).
Time-lapse sequences from 13-somite Mlc2a-/- embryos
showed that forward flow of erythroblasts was impaired. Erythroblasts were
either stationary (data not shown) or moving in an oscillatory pattern with an
equal amount of retrograde and anterograde motion
(Fig. 5 and see Movie 2 in the
supplementary material). Thus, instead of the rapid, pulsatile laminar-flow
patterns seen in wild-type embryos, slow oscillatory flows were observed in
Mlc2a-/- embryos. To determine how early the circulation is compromised, we examined plasma flow in Mlc2a-/- embryos using FRAP. Perfusion coefficients were comparable between wild-type (Fig. 3) and mutant littermates at 3- to 6-somites (data not shown), but by 8- to 9-somites, Mlc2a-/- embryos showed reduced plasma flow (Fig. 6). At 8- to 9-somites, we measured perfusion coefficients as high as 3835 µm2/second in wild-type embryos (24 measurements in 8 embryos) and often encountered regions with flow rates too fast to measure. By contrast, perfusion coefficients in mutant embryos were nearly four-fold lower (below 1045 µm2/second; 27 measurements in 9 embryos) and no vessels exhibited flow outside the measurable range. Of note, the perfusion coefficients measured in Mlc2a-/- embryos were still larger than pure diffusion (51±21 µm2/second), indicating that plasma flow was present, but weaker than in wild-type embryos. These data show that poor cardiac function can be detected in very early mutant embryos, soon after the heart begins to beat and before other phenotypes are evident.
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In control and untreated embryos, a hierarchical vascular branching pattern with enlarged avascular spaces was observed in yolk sacs after 24 hours in culture (Fig. 7A-C). Erythroblasts were robust in number and uniformly distributed within the vascular space (Fig. 7C). When erythroblasts were prevented from leaving the blood islands, vascular remodeling did not take place and the immature plexus persisted (Fig. 7D-F). Turning was also impaired in low-hematocrit embryos (Fig. 7D-F). To quantify these results, we established a scoring system for the extent of remodeling and turning (Table 2). These data confirm that reducing the hematocrit had a significant impact on yolk sac size, embryo turning and vessel remodeling. The methods used to reduce the hematocrit in embryos had no effect on plasma flow. Plasma flow rates were comparable to normal, untreated embryos (Fig. 8). Thus, we conclude that the circulation of plasma alone is insufficient to drive vascular remodeling, arguing against the idea that vascular remodeling is triggered by a soluble factor distributed by blood flow.
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), we
tested whether artificially increasing the viscosity of the erythroblast-free
plasma could restore remodeling. High-molecular-weight synthetic sugars
derived from plants, such as hetastarch, increase the viscosity of solutions
(Akers and Haidekker, 2004;
Haidekker et al., 2002). We
injected a hetastarch solution into the circulation of embryos with
sequestered blood cells and these embryos displayed clearly visible, large,
branched yolk sac vessels (Fig.
7G-I). These embryos were indistinguishable from wild type in
terms of the size of the yolk sac, and the embryo turning frequency was
restored (Table 2). Thus,
increasing the viscosity was sufficient to rescue the remodeling deficiency in
low-hematocrit embryos. We conclude that mechanical force, normally imparted
by the flow of circulating erythroblasts, is necessary and sufficient to
induce vessel remodeling in the yolk sac of early mouse embryos.
Expression of molecules involved in force transduction
Fluid-derived forces can directly influence cell-signaling pathways in
endothelial cells (see Li et al.,
2005). PECAM-1 and VE-cadherin (also known as Pecam1 and cadherin
5, respectively - Mouse Genome Informatics) are part of a complex that can act
as a mechanotransducer of shear stress
(Tzima et al., 2005). Also,
shear stress increases the levels of the mRNA encoding nitric oxide synthase 3
(eNOS; also known as Nos3 - Mouse Genome Informatics)
(Topper et al., 1996) and
induces changes in the subcellular localization of eNOS protein through a
PECAM-1-activated pathway (Cheng et al.,
2005; Dusserre et al.,
2004; Fleming et al.,
2005). To determine whether these molecules play a role in
force-related signaling in embryonic vessels, we examined the expression of
PECAM-1, VE-cadherin and eNOS in control, low-hematocrit and
low-hematocrit+hetastarch embryos. PECAM-1
(Fig. 9A-E) was observed on
endothelial and erythroblast cell membranes in both large
(Fig. 9A) and small
(Fig. 9B) vessels in control
embryos. PECAM-1 staining appeared somewhat reduced, but was still found on
the surface of endothelial cells of unremodeled vessels of the low-hematocrit
yolk sacs (Fig. 9C). Since
large vessels were not observed in the low-hematocrit vessels, only small
vessels are shown. Robust PECAM-1 staining was observed on the endothelial
cell membrane in both large (Fig.
9D) and small vessels (Fig.
9E) when plasma viscosity was restored. VE-cadherin expression was
also localized to the membrane of yolk sac endothelial cells and staining
appeared similar in all yolk sacs, indicating that the levels of VE-cadherin
protein are not dependent on fluid force
(Fig. 9F-J). In control
vessels, eNOS protein was found localized to the perinuclear region as well as
the cell membrane as has been described
(Cheng et al., 2005). eNOS was
barely detected above background in low-hematocrit embryos and localization of
eNOS to membrane or perinuclear regions was not observed. eNOS expression was
restored in low-hematocrit+ hetastarch embryos and membrane staining was
evident on large remodeled vessels, consistent with previous findings that
increased shear stress stimulates translocation of intracellular eNOS to the
cell membrane (Cheng et al.,
2005). Further, cell membrane proteins highlight the physical
changes of endothelial cells in response to changes in mechanical force.
Endothelial cells exposed to shear stress are known to elongate in the
direction of the flow (Dewey et al.,
1981), and elongated cells are evident in large vessels in
Fig. 9. Endothelial cells in
low-hematocrit vessels had a more cuboidal shape, but remained intact and
retained cell-cell associations. These results show that eNOS levels are
sensitive to changes in hemodynamic force during remodeling in the yolk sac
and that mechanosensory signaling pathways identified in vitro also appear to
regulate force responses in vivo during development
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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). Further,
defects in embryonic circulation in Mlc2a mutants could be detected
by FRAP before an obvious phenotype was discernable and before the steady flow
of erythroblasts is present. Thus, this method is effective in detecting
defects in cardiac contraction prior to the onset of remodeling and can be
helpful in determining whether remodeling defects are the secondary
consequence of impaired heart function.
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; Davies et al.,
1986; Levesque et al.,
1990), but the number of proliferating cells in the yolk sac is
similar in Mlc2a mutant and wild-type embryos (data not shown). Also,
culturing mouse embryos in KB-R7943, an Ncx1 inhibitor, results in impaired
remodeling, but does not induce oscillations (our unpublished data). Thus, the
remodeling defect in Mlc2a mutant embryos is likely to be due to
reduced laminar flow, rather than to the presence of abnormal oscillatory
flow.
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;
McGrath et al., 2003;
Phoon et al., 2000;
Phoon et al., 2002). By
physically blocking erythroblasts from leaving the blood islands, we have been
able to conclude that the onset of erythroblast circulation causes a change in
viscosity, which is necessary for vessel remodeling. This role for primitive
erythroblasts has not yet been described because many mutations that lead to a
reduction in primitive erythroblasts also affect endothelial cells directly
(Oike et al., 1999;
Oshima et al., 1996;
Robb et al., 1995;
Shalaby et al., 1995;
Shivdasani et al., 1995;
Suri et al., 1996;
Visvader et al., 1998). It is
assumed in many of these cases that disruption of the endothelial cell program
is the direct cause of the remodeling defect. We would argue that any
significant reduction in primitive erythroblasts alone could account for the
remodeling phenotype, but presently there is no way to eliminate the
erythroblast population exclusively. In addition to affecting vessel remodeling, reducing the hematocrit also resulted in impaired embryonic turning. Although it is not entirely clear why this occurs, yolk sacs of embryos with sequestered erythroblasts are slightly smaller than control embryos (Table 2). It is possible that poor circulation reduces the tension within the yolk sac or the embryo proper leading to impaired turning. Delays in turning were also observed in Mlc2a-deficient embryos (data not shown); however, we cannot assess whether this is a common feature of mice with hemodynamic deficiencies as it is generally not reported.
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) and chick (le Noble et
al., 2004) has shown that blood flow is required for the dynamic
remodeling of interconnections between vessels, but is not required for
vasculogenesis. Similarly, our data show that yolk sac vessels are formed and
can support flow by the time the heart begins to beat, but mature vessels do
not develop if viscous flow is not established. Consistent with this,
endothelial cells are found lining the yolk sac plexus by the time the heart
begins to beat (Ferkowicz et al.,
2003; Fraser et al.,
2005). Thus, it is clear that blood flow is not required for the
differentiation of endothelial cells.
Our work shows that the mechanical force exerted by blood flow imparts an
important signal to trigger vascular remodeling. Precisely which forces are
most important for vascular remodeling remains an open question, because both
shear stress and circumferential strain could be instructive. Shear stress is
directly dependant on the apparent viscosity of the fluid and is likely to be
the critical force acting on early endothelial cells during vascular
remodeling in the mouse embryo. Shear stress has been shown to induce changes
in gene expression, cytoskeletal architecture and proliferation in cultured
endothelial cells (Li et al.,
2005) and our previous work has shown that shear stress in early
mouse circulation is well within the range known to induce these types of
changes (Jones et al., 2004).
Despite our evidence supporting a role for shear stress, we cannot state that
shear stress is the only essential force at work in this system because
circumferential strain also contributes to forces felt by endothelial cells,
and we cannot distinguish a separate role for these types of forces during
embryogenesis.
Recently, there have been great strides in identifying the molecular
components of a shear stress transduction pathway, implicating many molecules
that are expressed during vasculogenesis and vessel remodeling
(Garcia-Cardena et al., 2001;
Li et al., 2005). Shear stress
is thought to be sensed by a functional complex of PECAM-1, VE-cadherin and
Vegfr2 (also known as Flk1 and Kdr - Mouse Genome Informatics) which, when
activated, in turn activates eNOS, PI3k (Pik3r1), chicken Src and the
conversion of integrins to high-affinity forms
(Fleming et al., 2005;
Orr et al., 2006;
Topper et al., 1996;
Tzima et al., 2005). Mutant
analysis has gleaned only limited information about the mechanisms of force
transduction in embryonic vessel remodeling. Surprisingly, PECAM-1-knockout
mice on two different background strains are viable to adulthood
(Duncan et al., 1999;
Schenkel et al., 2006). Vegfr2
plays a role in the early differentiation of mesodermal lineages
(Shalaby et al., 1997;
Shalaby et al., 1995) making
it difficult to assess later roles. VE-cadherin-deficient embryos fail to
remodel the initial vascular plexus
(Carmeliet et al., 1999;
Gory-Faure et al., 1999),
which would be predicted if endothelial cells could not properly sense shear
stress; however, endothelial cells become detached and undergo apoptosis,
which might be the primary defect
(Carmeliet et al., 1999).
Septal defects, deficiencies in cardiac maturation and function, vascular
dysfunction, reduced angiogenesis and increased mortality are hallmarks of
neonatal mice deficient in eNOS (Feng et
al., 2002; Lepic et al.,
2006; Zhao et al.,
2002), despite the fact that these mice can survive to adulthood
and reproduce. Thus, the involvement of these factors in mechanotransduction
in the embryo has been difficult to define through mutant analysis. Our data
show that eNOS is regulated by hemodynamic force in embryos, as would be
predicted by experiments showing that mRNA levels and the subcellular
localization of eNOS are regulated by shear stress in vitro
(Cheng et al., 2005;
Dekker et al., 2005;
Dusserre et al., 2004;
Fleming et al., 2005;
Topper et al., 1996). Also,
PECAM-1 and VE-cadherin persist on the cell surface in low-hematocrit embryos,
showing that endothelial cells remain intact and continue to express molecules
thought to be required for force activation. Continued expression of these
proteins would be expected because low-hematocrit embryos can be rescued by
restoring hemodynamic force. Overall, our data show that the mechanosensory
signaling pathways identified in cultured cells also appear to be functional
in vivo during development.
|
Conclusion |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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Supplementary material |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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|
ACKNOWLEDGMENTS |
|---|
-globin-GFP) mice and for helpful
discussions, Karen Hirschi and members of the Hirschi group for reading the
manuscript and for helpful discussions and Josephine Enciso for the PECAM-1
antibody. This work was supported by R01 HL HL077187 (M.E.D.), a T32 HL7676
Training Grant supporting J.L.L. and an AHA Graduate Fellowship for
E.A.V.J. |
Footnotes |
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REFERENCES |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONConclusionSupplementary materialREFERENCES |
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