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1 Department of Bioengineering, 2 Department of Medicine, and 3 Department of Orthopedics, University of California-San Diego, La Jolla, California 92093-0412; and 4 Department of Cell Biology, Duke University, Durham, North Carolina 27710
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Tropomodulins are a
family of proteins that cap the slow-growing end of actin filaments.
Erythrocyte tropomodulin (E-Tmod) stabilizes short actin protofilaments
in erythrocytes and caps longer sarcomeric actin filaments in striated
muscles. We report the knockin of the 
-galactosidase gene
(LacZ) under the control of the endogenous E-Tmod
promoter and the knockout of E-Tmod in mouse embryonic stem
cells. E-Tmod
/ embryos die around embryonic
day 10 and exhibit a noncontractile heart tube with
disorganized myofibrils and underdevelopment of the right ventricle,
accumulation of mechanically weakened primitive erythroid cells in the
yolk sac, and failure of primary capillary plexuses to remodel into
vitelline vessels, all required to establish blood circulation between
the yolk sac and the embryo proper. We propose a hemodynamic "plexus
channel selection" mechanism as the basis for vitelline vascular
remodeling. The defects in cardiac contractility, vitelline
circulation, and hematopoiesis reflect an essential role for E-Tmod
capping of the actin filaments in both assembly of cardiac sarcomeres
and of the membrane skeleton in erythroid cells that is not compensated
for by other proteins.
erythrocyte tropomodulin; cardiomorphogenesis; hematopoiesis; LacZ; yolk sac; vasculogenesis
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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ACTIN IS AN ABUNDANT PROTEIN in eukaryotic cells, where actin filaments form one of the three major cytoskeletal networks. Actin networks carry out cellular functions such as contraction, adhesion, and migration, which are important for normal physiology and embryonic development. In sarcomeres, six actin (thin) filaments interdigitate with each myosin (thick) filament, and, in other cases, actin filaments are organized into arrays such as contractile bundles, gellike networks, or tight parallel bundles. The polarized actin filaments undergo polymerization and depolymerization at both ends (38, 47, 61). The fast-growing (barbed) end polymerizes at up to 10 times the rate of the slow-growing (pointed) end, suggesting that the fast-growing end was more dynamic and important in assembly. The dynamics of actin networks are regulated by a number of actin-associated proteins, including those that cap either end of the filaments. The fast-growing end-capping proteins include adducin (25, 26) and CapG (33, 48). Interestingly, their null mutations are not lethal, only causing spherocytosis in erythrocytes (17) and impaired phagocytosis in macrophages (63), respectively. Knockout of gelsolin, which encodes a fast-growing end-capping and severing protein, also is not lethal (63). It is not known whether mice with targeted disruption of genes encoding slow-growing end-capping proteins would survive.
Tropomodulin was first isolated as a tropomyosin (TM)-binding protein (13) and later shown to be the long sought after slow-growing end-capping protein of the actin filaments (15, 18, 59). It increases the critical concentration of actin filaments, whose fast-growing end was capped by gelsolin, by converting ADP.P(i)-actin to ADP-actin at all slow-growing ends (60). The first tropomodulin identified was human erythrocyte tropomodulin (E-Tmod) (13), and several other tropomodulins have since been characterized. E-Tmod is highly conserved among humans (50), mice (19), rats (58), and chickens (3). E-Tmod is expressed not only in erythrocytes, but also in other tissues including lens, muscle, and embryonic stem (ES) cells (2, 8, 19, 64).
Several E-Tmod homologs encoded by distinct genes exist in humans, mice (10), and other species. These include N-Tmod in the rat brain (58), Sk-Tmod in chicken fast skeletal muscle (1), and ubiquitously expressed Tmod (U-Tmod) in various tissues (10). In mice, the identity between the amino acid sequences of E-Tmod and Sk-, N-, and U-Tmod is 56%, 57%, and 59%, respectively. Leiomodin in human extraocular muscles, sanpodo in Drosophila, and Tmod-like proteins in Caenorhabditis elegans also belong to this family (9, 12, 37, 62).
Tropomodulins may be targeted to different locations in cells to serve different functions. In chicken fast skeletal muscle fibers, the predominant Sk-Tmod is present at the pointed (free) end of the thin actin filaments in sarcomeres, whereas E-Tmod is recruited to costameric subsarcolemmal domains of the same cells (1). In human erythrocytes, E-Tmod is located at the junctional complex, which is the center of hexagonal lattices in the actin-spectrin-based membrane skeletal network (56).
E-Tmod binds to one end of the rodlike erythrocyte TM (14), specifically, to the NH2-terminal end of TM5 between residues 7 and 14 (52, 57). The complex formed by E-Tmod and TM5/5b functions as a molecular ruler for actin protofilaments (51). Protofilaments, uniformly 37 nm long (44), in turn define the hexagonal geometry of spectrin lattices in the membrane skeletal network, which supports the mechanical stability of the lipid bilayer and provides the elastic deformability of erythrocytes (51).
Several experiments have demonstrated a role for tropomodulin in striated muscle. In isolated skeletal myofibrils, antibodies made against E-Tmod were localized to the free end of the thin (actin) filaments in sarcomeres (15). In cultured embryonic chick cardiac myocytes, microinjected antibodies against E-Tmod led to marked elongation of actin filaments and reduction in the number of beating cells (18). Long actin filament bundles also formed when antisense E-Tmod mRNA was induced in fetal cardiomyocytes in culture (53) and peripheral myofibrils were disordered and lacked Z-lines. Conversely, overexpressing E-Tmod in cultured cardiomyocytes resulted in shorter thin filaments (28, 53), whereas overexpression in the hearts of transgenic mice led to dilated cardiomyopathy (54).
To reveal the consequences in mammals of a knockout of
E-Tmod on embryonic viability and development, particularly
in erythroid cells, which have never been reported, and the heart, we
disrupted the E-Tmod gene in mouse ES cells and obtained
mice that were null for the protein. We also knocked in a
-galactosidase reporter gene (LacZ) under the control of
the endogenous E-Tmod promoter that allows visualization of
those cells and tissues normally expressing E-Tmod, even in the absence
of the protein. E-Tmod/ embryos die around
embryonic (E) day 10 (E10) due to abnormalities in cardiac
contraction, vascular morphogenesis, and hematopoiesis. The lethality
of the E-Tmod/ mutation demonstrates the
importance of the slow-growing end of actin filaments in cellular
functions, and that the capping by E-Tmod during embryonic development
is not replaced by other proteins, including members of the Tmod family.
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EXPERIMENTAL PROCEDURES |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Construction of E-Tmod targeting vector.
Established gene targeting protocols were followed (20). A
1.7-kb 5' genomic fragment and a 5-kb 3' fragment of E-Tmod
flanking exon 1 (8), isolated from a mouse 129/Svj genomic
library (Stratagene; La Jolla, CA), were linked to a
LacZ-PGKneo cassette (6) at 5' (XhoI
site) and 3' (BglII site) positions, respectively, to construct the targeting vector (Fig.
1A). A thymidine
kinase (tk) gene was also linked to the 3' end of
the targeting cassette. In Fig. 1A, the tk gene
in the targeting vector is in gray and downstream of the 3' end of the
homologous region in black. The purpose of the tk gene was
to allow for negative selection to eliminate clones that had acquired
neomycin resistance (NeoR) through nonhomologous recombination.
However, because the positive selection by G418 had allowed us to
obtain two E-Tmod+/ ES cell clones through
homologous recombination, negative selection was never implemented.
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Generation of E-Tmod knockout mice.
The linearized targeting construct was electroporated into cultured ES
cells (R1) (30), and G418-resistant colonies were screened
for homologous recombination by PCR and Southern blot analysis. The
Tmod+/ ES clones expanded in culture were
injected into normal blastocysts of C57/black, 3 days after their
fertilization, to create chimeric mice. The interbreeding of
Tmod+/ mice was used to generate
Tmod/ mutants.
Genotyping of E-Tmod knockout ES cells and mice. Genomic DNA was isolated from ES cells, yolk sacs, or tails from the embryo proper or mice (DNeasy Tissue Kit, QIGen). Genotypes were examined first by PCR with a 5' primer screen-P1 (5'-ATGCTCCTGGGTGACTAAGGTG-3') and a 3' primer mTmod140R (5'-CAGCTCCTCCTCTGTGAGG-3') for wild type or a 3' primer Lac541R (5'-CAGGTCAAATTCAGACGGCA-3') for the disrupted gene. Samples containing disrupted genes were confirmed by Southern hybridization with both 5' and 3' probes marked in Fig. 1A.
Phenotype analyses. Whole mount LacZ staining, in situ hybridization, and immunohistological analysis were performed according to established methods (29, 42, 65), respectively.
Transmission electron microscopy.
E9.5 wild-type and E-Tmod/ embryos were
immersed in a glutaraldehyde-tannic acid fixative in buffered mammalian
Ringer solution composed of (in mM) 150 NaCl, 5 KCl, 1.5 CaCl2, 1.5 MgCl2, and 10 3-(N-morpholino)propanesulfonic acid (pH 7.0). It was
followed by osmium tetraoxide secondary fixation and block stained in
uranyl acetate. Thin sections of the araldite-embedded blocks were
stained with KMnO4/Pb, as described by Nassar et al.
(31).
Cytospin. A cytocentrifuge (Cytospin 2, Thermo Shandon; Pittsburgh, PA) was used to spin (at 28 g) mouse primitive erythroid cells collected from yolk sacs.
Micropipette aspiration technique. The micropipette aspiration technique has been reported and used extensively to characterize mechanical properties of individual human erythrocytes, white blood cells, and nuclei (5, 11, 49). Deformation tests for primitive reticulocytes and erythroblasts were performed with modifications for their larger sizes and/or the presence of the cytoskeleton.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Generation of E-Tmod knockout mice. A targeting vector was constructed by replacing exon 1 of mouse E-Tmod, which contains the translation initiation codon (8), with bacterial LacZ and NeoR genes (Fig. 1A).
Targeted disruption of E-Tmod was accomplished by homologous recombination in mouse ES cells (20). After electroporation and G418 selection, 288 surviving ES cell colonies were genotyped by PCR screening, followed by Southern blot analysis (Fig. 1B). In PCR, the wild-type gene generated a 1.7-kb fragment, whereas the disrupted gene generated a 2.2-kb fragment. In Southern blot analysis, the 5' probe (position shown in Fig. 1A) hybridized to a 4.7-kb fragment in the wild-type gene and a 7.2-kb fragment in the disrupted gene (Fig. 1B). Two independent E-Tmod+/ ES cell clones, A20
and B9, were established. Microinjecting
E-Tmod+/ ES cells into normal blastocysts
created seven male chimeric mice. Two (both derived from A20) had a
germ line transmission of the disrupted E-Tmod gene and gave
rise to several E-Tmod+/ mice,
which survived to adulthood and were fertile.
Earlier in culture, G418 was increased from 0.25 to 4 and 6 mg/ml to
establish two E-Tmod/ ES cell
lines. Although E-Tmod/ ES cells were not
chosen in creating chimeric mice because of the concern that they may
not contribute to the germ line, their establishment proved the
viability of E-Tmod/ ES cells.
E-Tmod null mutation is embryonically lethal.
Screening of 177 offspring produced by intercrossing
E-Tmod+/ mice revealed no
E-Tmod/ littermates, suggesting that an
E-Tmod null mutation is embryonically lethal.
Because E-Tmod/ ES cells are viable, we
investigated the timing of lethality by genotyping embryos from E8.5 to
E13. Whereas wild-type, E-Tmod+/, and
E-Tmod/ embryos exhibited ~1:2:1 ratios,
respectively, in the early stages, no
E-Tmod/ embryos were found alive after
E10.5, indicating that they had died ~E10.
Complete block of E-Tmod protein synthesis in
E-Tmod/
embryos.
Western blot analysis with the use of E-Tmod monoclonal antibody 204 (Fig. 1C) revealed that, whereas
E-Tmod+/ embryos had about one-half of E-Tmod
protein present in the wild type, E-Tmod/
embryos had no detectable E-Tmod protein. Note that only exon 1 and
small portion (0.8 kb) of intron 1 were replaced and the resulted
sequence was confirmed by sequencing the PCR fragment derived from this
mutated region. Because no upstream promoter sequence is affected,
cells are able to transcribe the 5' untranslated region of
E-Tmod (8) and the LacZ sequence,
which has a stop codon. Furthermore, the downstream E-Tmod
sequence is out of the correct reading frame after the insertional mutation.
E-Tmod expression reported by knocked-in bacterial LacZ gene.
-Galactosidase, the gene product of LacZ, generates blue
signals by hydrolyzing
5-bromo-4-chloro-3-indolyl--D-galactopyranoside (X-Gal).
Therefore, the knocked-in LacZ under the control of the endogenous E-Tmod promoter (Fig. 1A) signals the
pattern of expression of E-Tmod protein in heterozygotes as well as the
promoter activity of E-Tmod in
E-Tmod/ cells, even in the absence of the
expression of E-Tmod protein.
embryos (Fig. 1D), the
whole mount X-Gal staining (29) revealed no detectable
LacZ expression at E7.5. A high-level expression of
LacZ was first detected at E8.0 in blood islands or primary
capillary plexuses in the yolk sac and the developing heart tube in the
embryo proper. The expression of LacZ revealed the detailed
organization of primary capillary plexuses and fused linear heart
tubes, from the developing atria (two arches at the bottom of the heart
tube) to fused ventricles. The normal rightward looping of the heart
tube was obvious in the E-Tmod+/ embryo at
E8.5. Later, at E9.5, the heart of heterozygotes continued to express
lacZ, whereas no detectable signals were found in developing somites (20-29 somite pairs stage). At E9.5 in heterozygotes, circulating LacZ-expressing blood cells also marked dorsal
intersegmental (or intersomite) arteries, anterior cardinal veins, and
other blood vessels in the embryo proper. In
E-Tmod/ embryos, even in the absence of
E-Tmod protein, the expressed LacZ reporter gene under the
control of the E-Tmod promoter also highlighted the
"null" tissues that normally express E-Tmod. The blue staining
facilitated the analyses of disease phenotypes in cardiomorphogenesis,
vascular morphogenesis, and hematopoiesis in
E-Tmod/ embryos.
Disease phenotype in heart.
We examined >50 E-Tmod/ embryos for the
null phenotype presented. The E-Tmod/ embryo
was grossly normal at E8.5. Thereafter, significant growth retardation
was observed. Figure 2a shows
left views of a wild-type embryo and an
E-Tmod/ embryo at E9.5. The abnormal heart
with a massive pericardial effusion in the
E-Tmod/ embryo is apparent and consistent
with embryonic heart failure. The decreased growth of the facial and
brain primordial is also noticeable, which may be caused by the
complete deficiency of E-Tmod or the secondary effect resulting from
the lack of circulation into those regions.
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Contractility of E-Tmod/ heart
tube.
When E9.5 embryos were removed from the uterus, the developing hearts
of wild-type embryos had peristaltic contractions (Fig. 2a,
top). In contrast, those of
E-Tmod/ embryos had barely noticeable
tremors, initiated from the region corresponding to atria (Fig.
2a, bottom).
heart tube at E9.5 looped normally,
with atria, ventricular chambers, and an outflow tract in place. But
E9.5 E-Tmod/ mutant had a single, fused
heart chamber with overdeveloped trabeculae. A higher magnification
reveals individual cardiomyocytes expressing LacZ (Fig.
2c). The trabecular myocytes were increased and more discontinuous in the Tmod/ mutants. It would
be of interest to investigate why this is the case.
Looping of
E-Tmod/
heart tube.
The E-Tmod/ heart tube was distorted after
initial looping at E8.5. Sequential front views of X-Gal-stained heart
tubes of E-Tmod+/ (Fig. 2d) and
E-Tmod/ (Fig. 2e) embryos were
compared. Between E9.0 and E10.5, E-Tmod/
embryos showed abnormal looping. Without the outgrowth of the right
ventricle, the single ventricle was abruptly connected to the outflow tract.
Expression of cardiac-specific markers.
To assess the degree of development and specification of the
E-Tmod/ heart, four cardiac-specific
markers, myosin light chain (MLC)2a, MLC2v, dHAND, and eHAND, were used in
whole mount in situ hybridization. E-Tmod/
embryos at E9.0 (Fig. 2, f-i,
bottom), which began to exhibit retarded growth, were
compared with wild-type embryos (Fig. 2, f-i,
top). The images have the same right side views except for Fig. 2i, which has a left-front side view for left ventricles.
/ embryo, MLC2a was
expressed in the entire heart (Fig. 2f, bottom), with a near-normal expression level, although the entire heart was
slightly smaller. MLC2v (Fig. 2g,
bottom) showed a restricted expression, indicating that
atrioventricular chambers were specified in the
E-Tmod/ heart tube. But the area of
MLC2v expression was much smaller compared with the
wild-type heart tube (Fig. 2g, top), suggesting that perhaps only one ventricular chamber was developed.
To further assess the development of right and left ventricular
chambers in the E-Tmod/ heart, the
expression of two transcription factor genes was analyzed. Normally the
expression of dHAND (46) and eHAND
(39) is predominantly restricted to the region of the
looping heart tube that gives rise to the right and left
ventricle, respectively. The lack of dHAND expression (Fig.
2h) is concomitant with the single ventricular chamber
expression pattern of MLC2v (Fig. 2g) and is
consistent with the lack of right ventricle in
E-Tmod/ mutants. Although the expression of
eHAND was detectable, it appeared downregulated in the
E-Tmod/ heart tube (Fig. 2i),
suggesting that even the left ventricle was not fully specified.
Transmission electron microscopy analysis of
E-Tmod/
cardiomyocytes.
To assess whether the abnormal looping and the lack of contraction of
the E-Tmod/ heart reflected abnormal
organization of actin filaments in cardiomyocytes in the absence of
E-Tmod, we examined thin sections (30-40 nm) of wild-type and
E-Tmod/ embryonic hearts by transmission
electron microscopy. At E9.5, sarcomeric myofibrils had been organized
in the wild type (Fig. 3A)
with normal M lines and Z lines. Most of the actin filaments were
already interdigitated with thick filaments and incorporated into
regular sarcomeres. Filaments had normal and uniform lengths and
attached to the membrane at nascent intercalated disk junctions (Fig.
3E).
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/ cardiomyocytes, in the most
ordered areas (Fig. 3B), filaments were gathered into
myofibril bundles with Z lines being replaced by elongated, stress
fiberlike dense bodies (Zdb). In many areas, numerous thin and thick
filaments formed very disordered arrays with amorphous dense Z bodies
scattered around them; only a few thin and thick filaments were
interdigitated (Fig. 3C). Sometimes the thin and thick
filaments were segregated into separate bundles (Fig. 3B),
which is rare in wild-type embryos. Intercalated disks and gap
junctions were also developing in E-Tmod/
cardiomyocytes. However, more dense material was associated with the
nascent intercalated disks in the E-Tmod/
cells compared with that of wild-type embryos (Fig. 3E).
Disease phenotype in vascular morphogenesis in yolk sacs.
The primary capillary plexuses in yolk sacs were first revealed at E8.0
in E-Tmod+/ embryos (Fig. 1D) by
LacZ-expressing erythroblasts (see below). Before then, at
E7.5 when blood islands normally form, there was no detectable
LacZ expression to reveal the structure of blood islands. At 8.5, E-Tmod+/ and
E-Tmod/ yolk sacs had similar organization
of primary capillary plexuses (Fig.
4a). The heart tube was also
visible, which should have initiated the rightward looping (Fig. 2,
d and e), normally been beating
(40), and connected to primary capillary plexuses as normal vitelline circulation starts at E8.5 (16, 21). At
E9.0, the E-Tmod+/ yolk sac had noticeable
remodeling of blood vessels (Fig. 4b). The process appeared
to have direction because all developing vessels radiated from the
largest vessel, and the diameter of each vessel tapered from large to
small. At E9.5, mature vitelline vessels (Fig. 4c) had been
formed.
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/ mutants, the remodeling of the
primary capillary plexuses into treelike mature blood vessels was
arrested (Fig. 4, bottom). At E9.0, primary capillary
plexuses grew to occupy a larger area but remained a honeycomb-like
network of channels (Fig. 4b, bottom). These
plexuses fused around E9.5, forming large blood pockets (Fig.
4c, bottom). Cross sections of the X-Gal-stained yolk sac of E-Tmod/ (Fig. 4d)
revealed packed primitive erythroblasts expressing high levels of
lacZ. Therefore, it is the blood cells that outlined primary
capillary plexuses and blood vessels when stained by X-Gal.
Disease phenotype in hematopoiesis.
Normally, the hematopoietic activity in mice shifts from the yolk sac
(primitive lineages) to the developing fetal liver (definitive lineages) between E10 and E12, with the seeding of hematopoietic stem/precursor cells derived from the yolk sac and/or the
pleurosplanchic (P-Sp)/aorta-gonads-mesonephros region (7, 34,
22). At E9.5, wild-type and
E-Tmod+/ embryos had a large number of blood
cells circulating in the cardiac chambers, dorsal aorta, and blood
vessels in both the embryo proper and yolk sac (Fig.
5a-d, top, and Fig.
1D). The insets in
Fig. 5b, top, also show some "blue"
cells outside of chambers. The E-Tmod/
embryo proper, however, was "anemic" (Fig. 5a and
b, bottom). Often, the entire transverse sections
of an X-Gal-stained embryo proper (Fig. 5b,
bottom) revealed no visible blue blood cells. In one case,
some blood cells were found inside the atria (Fig. 2b,
bottom), and in another case, a small pink region was
noticed near the heart chambers of an unstained embryo (Fig.
5a, bottom). The anemia suggests limited
hematopoiesis in the E-Tmod/ embryo proper,
lack of vitelline circulation, and/or a defect in primitive blood cells
in the yolk sac. The cross sections of the embryo proper also revealed
that vascular chambers in the E-Tmod/ embryo
proper appeared to be lacking or minimized compared with the normal
(Fig. 5b).
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Light microscopy of blood cells from yolk sacs.
Blood cells from E9.5 yolk sacs were examined under light microscopy
and several abnormalities were found in
E-Tmod/ cells. Figure 5e,
left, for wild-type cells shows (from left to
right) one primitive reticulocyte or erythrocyte, one
erythroid cell undergoing enucleation, and one erythroblast. Their
large size (~29 µm in diameter) indicated the primitive origin,
because a definitive erythrocyte in adult mice was only ~6 µm.
Figure 5e, right, for
E-Tmod/ cells shows (from left to
right) one of very few primitive reticulocytes or
erythrocytes, one binucleated primitive erythroblast, and one nucleated
erythroblasts that had a membrane hernia. Often,
E-Tmod/ erythroid cells were partially
hemolyzed. Binucleated erythroblasts were also found in cross sections
of yolk sacs (Fig. 5d, bottom) and on cytospin
slides (see below; Fig. 5f, right). In fact,
~2% of the primitive blood cells examined were binucleated (5 of
246), whereas none was observed in the similar cell count of the wild type. This finding suggests that cytokinesis in E-Tmod null
mutants may either be slowed or sometimes incomplete.
Cytospin of yolk sac blood cells.
Blood cells collected from yolk sacs were further examined by clinical
cytocentrifuge, followed by Giemsa staining.
E-Tmod+/ blood cells (Fig. 5f,
left) were flattened on the slides but remained intact. At
E9.5, most of the cells were differentiating erythroblasts, as reported
by Palis and Yoder (35). One was seen in the process of
enucleating, ~3 days earlier than E13, as reported by Bethlenfalvay
and Block (4). In contrast, no E-Tmod/ cell membranes or cytoplasm remained
associated with the nuclei of erythroblasts. The cytospin (Fig.
5f, right) captured several erythroblast nuclei,
a binuclear remnant, and a broken membrane away from the cell body. All
of these findings suggest that mechanical strength of
E-Tmod/ erythroid cells is weakened.
Mechanical strength of blood cells tested by micropipette
aspiration.
To test the mechanical strength of E-Tmod/
erythroid cells, membrane deformation and fragmentation analyses were
performed with the micropipette aspiration technique (5,
49). Primitive erythroid cells retain nuclei during most of
their lifespan and at E9.5; only very few enucleated cells were
found from the yolk sac. The membrane deformation under a microscope in
response to a step-negative pressure applied to a micropipette was
recorded (Fig. 5g). In addition, membrane fragmentation
induced with a higher negative pressure and the stress at which
fragmentation occurred was also calculated based on geometry and the
applied pressure (Fig. 5h). Figure 5g,
top, shows a deformed length of a wild-type enucleated
erythrocyte (or reticulocyte) membrane, which was about four times the
pipette radius (Rp), and Fig. 5g, bottom, shows the deformation length of an
E-Tmod/ erythrocyte membrane (or
reticulocyte), which was about eight times the
Rp under the same condition. This finding
suggests an overextension of the membrane skeleton without E-Tmod in
response to a given negative pressure.
1.
The work is presented as the product of aspiration force (in dyn/cm)
and aspiration duration (in seconds) when the membrane started to
fragment. At this level of work, 45% of 31 wild-type cells tested had
membrane fragmentation. Fig. 5h, bottom, shows the response of a nucleated erythroblast from an E9.5
E-Tmod/ yolk sac under the same condition.
Arrow 1 was not labeled because the membrane nodule had been
aspirated into the pipette, and the work for fragmentation was 11.25 dyn · s · cm1.
At this level of work, 64% of 36 E-Tmod/
erythroblasts tested had membrane fragmentation. Therefore, both deformation and fragmentation analyses indicated that
E-Tmod/ erythroid cell membranes had less
mechanical strength than those of the wild type. The erythroid cells
without E-Tmod are easier to overextend (by ~2×) and to form nodules
(by ~4×), leading to partial hemolysis and fragmentation in response
to mechanical stress.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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The E-Tmod knockout and LacZ knockin
strategy provides the expression pattern of E-Tmod with
single cell resolution and disease phenotypes in null mutants without
the concerns of cross hybridization and cross reactivity. The
cell-specific expression pattern of E-Tmod reported by
LacZ agreed with the tissue-specific expression patterns
detected by in situ hybridization (19), supporting the
assumption that the knocked-in LacZ expression pattern in E-Tmod+/ embryos reflected the endogenous
E-Tmod expression pattern. The only exception was somites at
E9.5, where the expression of the knocked-in LacZ was not
detectable, but a very weak signal was detected by in situ
hybridization. Three possibilities for the discrepancy are the
following: 1) the disruption of a portion of the endogenous
promoter for E-Tmod required for the expression in somites
or skeletal muscle; 2) the different sensitivities between
these two techniques; or 3) the cross hybridization of the
E-Tmod cDNA probe to transcripts of E-Tmod
homologs. It is known that Sk-Tmod is expressed in certain
types of skeletal muscles and U-Tmod is expressed
ubiquitously, at least in adult tissues. To minimize the disruption of
endogenous promoters, no sequence upstream from exon 1 was disrupted in
our design. Intron 0, which is upstream from exon 1, and exon 0, which
is further upstream and potentially contains the transcription
initiation site (8), were all intact, with their sequences
confirmed. Only exon 1, which does not contain the transcription
initiation site but does contain the translation initiation codon ATG,
and a small portion of the downstream intron 1 (0.8 kb), were replaced
by LacZ. Therefore, unless there were promoter elements
specific for somites that are located within exon 1 or the small region
of intron 1, the LacZ expression pattern may represent that
of the endogenous E-Tmod. We are in the process of examining
the spatial-temporal expression pattern of LacZ later than
E9.5, to determine the exact timing of LacZ expression in
somites (and other tissues) because a strong LacZ expression
has been detected in skeletal muscles in the adult (data not shown).
Whether cells from E-Tmod+/ heterozygotes are
totally equal to those of true wild types raises an interesting
question, and we are systematically examining tissues/cells in adult
E-Tmod+/ mice and comparing them with
wild-type mice. Our preliminary results on the definitive erythrocyte
deformation by micropipette aspiration suggest no significant
difference between E-Tmod+/ heterozygotes and
wild-type cells (C. Vera, K.-L. P. Sung, and L. A. Sung, unpublished results).
In the complete absence of E-Tmod capping of the slow-growing end of actin filaments, the processes of cardiac and vascular morphogenesis and hematopoiesis are arrested. These disease phenotypes in developing embryos may be better understood by discussing the role of E-Tmod in the erythrocyte membrane skeleton, where the molecular organization and mechanical properties have been characterized most extensively.
In definitive human erythrocytes, a complex formed by globular E-Tmod
and rodlike TM5/5b (33-35 nm long) caps and protects a short
protofilament of 37 nm (51, 57). Each of these short protofilaments is knitted by six spectrins to form a thin, continuous, elastic protein network that supports the lipid bilayer during deformation. Although the detailed organization of the membrane skeleton may not be identical in primitive and definitive erythroid cells, or before and after enucleation, the role for E-Tmod may be
similar. Without E-Tmod to cap short protofilaments, the network may
become thicker (some actin filaments may be longer than 37 nm), patchy
(some longer actin filaments may be bundled by >6 spectrins), and
discontinuous (some other actin filaments may not be connected to
neighboring actin filaments by spectrins). The remnants of
E-Tmod/ membrane skeletons collected by
cytospin fit the predicted discontinuity of the network. Such
disorganization results in a weaker tensile strength, allowing the
membrane skeletal network to overextend (hemolysis) and break
(fragmentation) in response to mechanical stress.
Primitive erythroblasts and cardiomyocytes are the two cell types that show upregulation of E-Tmod at E8.0, when they begin passive and active deformation, respectively. This finding agrees with the previous report by in situ hybridization, which located the E-Tmod gene transcript in blood islands and myocardium (19). The heart tube normally starts peristaltic contraction at E8.0 (40) and the onset of circulation is at E8.5 (16, 21). The upregulation of transcription probably coincides with a functional requirement for E-Tmod and may be mechanically regulated. Examples of genes whose promoter elements bind to transcription factors in response to mechanical signals, such as shear stress, have been reported (45, 67). Increasing E-Tmod protein concentration may ensure sufficient number of short protofilaments to build a membrane skeletal network with suitable mechanical strength in erythroblasts, enabling them to survive the deformation induced by cardiac contractions. In cardiomyocytes, E-Tmod may serve a dual role, capping and stabilizing both sarcomeric actin filaments and those associated with the membrane cytoskeleton, thereby conferring mechanical strength during contraction cycles.
The phenotype of blood cells and cardiomyocytes derived from
E-Tmod null ES cells differentiated in vitro is one of the
interesting projects we have proposed to do. ES cell-derived blood
cells and cardiomyocytes may be characterized and compared with those
derived in vivo. Along this line, we have been able to culture
contractile cardiomyocytes from normal and
E-Tmod+/ E9.5 embryos, but not from
E-Tmod null embryos (I. Lian, X. Chu, and L. A. Sung,
unpublished results).
The E-Tmod/ heart tube was not contractile
at E9.5, which is consistent with the lack of organized sarcomeres in
cardiomyocytes. It was less obvious why the myofilaments were in such
disarray in the absence of E-Tmod. In cultured chick cardiomyocytes,
E-Tmod was reported to function late in myofibrillogenesis, maintaining the final length of thin filaments after they have been assembled (18). Therefore, if thin filaments in
E-Tmod/ cardiac myocytes were organized in
sarcomeres, but longer than normal in the absence of E-Tmod, it would
be consistent with a late function in sarcomere assembly. In contrast,
in cultured chick skeletal myocytes, E-Tmod was reported to function
early, appearing at the earliest nonstriated myofibrils and capping
actin filaments even before the filaments are cross-linked into
Z-bodies by 
-actinin (1). Z-bodies and actin filament
bundles near the sarcolemma have been proposed to serve as scaffolds
for the development of mature sarcomeres (32, 43). If
E-Tmod functions early in mouse cardiac myofibrillogenesis, lack of
E-Tmod could interfere with the assembly of sarcomeres: actin filament
length cannot be regulated and Z-bands are not assembled. The scattered Z bodies and disordered myofilaments we observe in E-Tmod
null hearts may be the remnants of this disrupted process. Consistent with this idea is the "abnormal" coincidence of E-Tmod and
-actinin at Z-disks observed in cardiomyocytes of
E-Tmod-overexpressing transgenic mice (54).
Furthermore, in chicken skeletal muscles, E-Tmod and spectrin have been
colocalized to costameres overlying the Z-band with flanking I bands,
or I-Z-I bands (1). Costameres are riblike cytoskeletal
structures attached to the sarcolemma that broaden and narrow in
concert with the underlying I bands in contractile cycles
(36). If E-Tmod stabilizes the actin-spectrin based
membrane skeleton in costameres, it may allow the sarcolemma to sustain
deformation. Without E-Tmod, this membrane skeleton may be mechanically
weakened and cardiomyocytes may be damaged during contractions.
E-Tmod is not required for the initial rightward looping of the heart tube, but is needed for the outgrowth of right ventricle. This may be due to a potential role for E-Tmod in cell fate decisions or the consequence of heart failure. The former is suggested by the finding that sanpodo (a homolog of tropomodulin) is a lineage gene in Drosophila (37) and asymmetric cell divisions may play a major role in cardiac and somatic muscle patterning. The latter is suggested by the finding that the "diminished right ventricle" is shared by several null mutants of cardiac-specific transcription factors, such as MEF2C (27), dHAND (46, 66), and Csx/Nkx2.5 (55), and the Ncx1-null mouse models. Ncx1 encodes a sodium-calcium exchanger; the null mice lacked spontaneous heartbeats but showed normal cardiomorphogenesis (up to E11) except that the future right ventricle region was severely underdeveloped (23).
Several of these mutants (e.g., of dHAND, Csx/Nkx2.5, and Ncx1) also exhibited arrests in vascular morphogenesis. Vasculogenesis is the process by which primary capillary plexuses form channels of similar diameters in situ. Angiogenesis that follows is thought to involve multiple phases, including new channel formation (by sprouting and nonsprouting), segregation of arteries and veins, establishment of endothelial cell-cell junctions, and regression of vascular channels (see Ref. 41 for a review). The arrest of vascular remodeling after vasculogenesis concomitant with cardiac failure due to completely unrelated defects (transcription factors, Na/Ca exchanger, and structural protein) suggests that cardiac contraction-derived pressure gradient and blood flow may be critical. Previously, without real-time scale and single cell resolution, it has been difficult to document how the primary capillary plexuses remodel (41). Here, the LacZ reporting allows documentation of remodeling at the single cell level between E7.5 and E10, although not on a real-time scale.
We propose a plexus channel selection mechanism based on our
observation that the zigzag contours of forming blood vessels (Figs.
4b and 6a) resemble the geometry of and are
continuous with the channels of primary capillary plexuses (Fig.
4a). We define a "capillary channel" at this stage as
the space occupied by blood cells and not that defined by an
endothelium. Our hypothesis suggests that vitelline blood vessels form
by selecting some preexisting channels and abandoning some others
within the orthogonal network of channels, depending on their alignment
with the pressure gradient and flow established by the cardiac output
(Fig. 6b). With time, the
blood flow smoothes out sharp angles and the blood pressure increases
diameters of blood vessels (Fig. 6, c and d),
after endothelial junctions seal the vessels.
|
How are some capillary channels selected and others abandoned? Because
primary capillary plexuses formed in situ are remodeled into
endothelial cell-lined vessels with blood cells flowing inside, stationary blood cells grown in situ must become mobile. It is reasonable to assume that at this point, differentiated erythroblasts (stained blue in Fig. 6a, also see Fig. 5) have lost
adhesive contacts with each other, whereas other cells in "white
islands" (mesoderm/mesenchymal cells not expressing LacZ)
still adhere to each other. Initially, blood cells in capillary
channels near the outflow tube and those aligned approximately with the
pressure gradient generated by cardiac output (marked by long arrows)
would be pushed downstream by the positive pressure. These capillary channels would stay open and develop into mature blood vessels, thus
being selected. Others that connect selected channels are often
approximately perpendicular to the pressure gradient and may only
experience a relatively small pressure gradient between two ends. Blood
cells in these channels (flanked by short arrows in Fig. 6a)
may move slowly toward the two selected channels at the two opposite
ends. As the space left behind is taken up or overgrown by adjacent
nonblood cells, these capillary channels would disappear and thus be
abandoned (Fig. 6). At the other (future venous) end, the blood flow in
capillary channels toward contractile atria keeps them open, and they
subsequently develop into veins. E-Tmod/
embryos without effective peristaltic contraction of the heart may,
therefore, be unable to select and abandon channels, thus remodeling
primary capillary plexuses (a network of channels) into a mature
vasculature (a treelike organization of channels) that consists of
arteries, capillaries, and veins.
Figure 1D shows no X-Gal staining between the heart and yolk
sac in normal E8.0 embryos, suggesting little, if any, blood cells are
present in this region at this stage. At E8.5, the X-Gal staining has
become continuous between the heart and the capillary plexuses (at
least grossly) in both E-Tmod+/ heterozygotes
and null mutants (Fig. 4a), suggesting this region is
"patent" even in the null mutant. It is after E8.5 that the process
of conversion of a primary capillary plexus into blood vessels in the
E-Tmod null mutant is disrupted.
It is not clear from this study whether there are physical structures or endothelial cells in the capillary channels separating blood cells and surrounding mesodermal cells, like in the mature blood vessels. Our finding that flattened cells along the internal walls of mature vessels (data not shown) did not express detectable E-Tmod by X-Gal and that several other mutations have the same phenotype (i.e., fail to remodel), even though they do not affect vessel cells, further support the notion that pressure gradient-derived blood flow provides the mechanism to select capillary channels from a network and convert them into mature blood vessels.
Although we hypothesized that the arrest of vascular morphogenesis is
mainly due to the lack of pressure gradient across the field of primary
capillary plexuses because of the failure of cardiac contraction, we
have not excluded the possibility that the complete deficiency of
E-Tmod in endothelial cells and/or smooth muscle cells may also
contribute to the arrest. Immunocytochemistry using an endothelial cell
antibody (e.g., that against platelet endothelial cell adhesion
molecule) and anti-smooth muscle actin antibody on embryos between E7.5
and E10 would provide useful information. Our preliminary analysis on
E9.5 embryos showed that the signals for platelet endothelial cell
adhesion molecule in the wild type corresponded to the walls of the
developing vasculature, but no signals corresponding to the walls of
the primary capillary plexuses were found in the
E-Tmod/ yolk sac. The signals for smooth
muscle cell actin, however, were similar between wild-type and null
mutants (X. Chu and L. A. Sung, unpublished data). We have not
looked at the placenta to see whether the vasculature of the null
mutants is normal.
In conclusion, E-Tmod null mice whose actin filaments are
not capped by E-Tmod at the slow-growing end develop disease phenotypes in each of the three key components of the circulatory system: blood
cells, the heart, and blood vessels. The end result is lack of
circulation. Lacking circulation at this developing stage, embryos
would not survive. Lacking circulation between the embryo proper and
the yolk sac also makes E-Tmod/ embryos a
potential model to dissect the source of definitive hematopoietic
stem/precursor cells. Whether the P-Sp region of an
E-Tmod/ embryo displays definitive
hematopoietic potential between E8.5 and E10, or whether yolk
sac-derived E-Tmod/ cells at E9.0 are
capable of reconstituting the hematopoietic system in newborn mice
(68), is of great interest. If
E-Tmod/ yolk sac/P-Sp stem cells do
reconstitute the hematopoietic system, there would be a large number of
definitive E-Tmod/ erythrocytes circulating
in wild-type mice. This would allow their membrane skeletal network to
be examined by electron microscopy and mechanical properties
characterized by micropipette aspiration. The role of E-Tmod in
cell membrane mechanics with the use of the simplest cell model would
then be established.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Dr. Kenneth Chien for making this investigation possible and Dr. Silvia Evans for suggesting cardiac chamber-specific transcription factor experiments and nucleotide probes. We thank Dr. Geert Schmid-Schönbein for the use of his microscope and discussion of blood vessel remodeling. We appreciate Drs. Chenleng Cai, Mark F. Couglin, and Stephen M. Baird for help with in situ hybridization, image recording, and cytospin, respectively. We also thank John Kim for assistance in micropipette experiments and Lynn Truong for manuscript preparation. We used the Molecular Biology Common Facility in the Department of Bioengineering established with the support by the Whitaker Foundation. We also utilized the Transgenic and Gene Targeting Core Facility in the Cancer Center at University of California-San Diego for gene targeting.
| |
FOOTNOTES |
|---|
This work was supported by National Heart, Lung, and Blood Institute Grant PO1HL-43026-6 and Training Grant HL-07089 (to X. Chu). C. Vera was supported by a University of California Institute for Mexico and the United States-Consejo Nacional de Ciencia y Tecnologia predoctoral fellowship.
Address for reprint requests and other correspondence: L. A. Sung, Dept. of Bioengineering and Center for Molecular Genetics, Mail Code 0412, Univ. of California-San Diego, 9500 Gilman Dr., La Jolla, CA 92093-0412 (E-mail: amysung{at}bioeng.ucsd.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published January 23, 2003;10.1152/ajpheart.00947.2002
Received 4 November 2002; accepted in final form 17 January 2003.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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P = 30,000 dyn/cm2; duration,
t = 1,300 ms). In the fragmentation test (h,
Rp = 0.75 µm, aspiration force = 22.5 dyn/cm), the cell membrane showed progressive deformation with a
nodule (arrow 1), a "neck" (narrowing membrane segment,
arrow 2), a cell body (arrow 3), and a partially
deformed nucleus membrane (arrow 4, not seen in wild type).
Bars = 0.5 mm for a-c and 20 µm for
d-f.
