1 Department of Molecular Cell Biology, Cardiovascular Research Institute Maastricht (CARIM), University of Maastricht, PO Box 616, 6200 MD Maastricht, The Netherlands
2 Department of Anatomy & Embryology, Molecular and Experimental Cardiology Group, Academic Medical Center University of Amsterdam, 1105 AZ Amsterdam, The Netherlands
3 Department of Biochemistry, CARIM, University of Maastricht, PO Box 616, 6200 MD Maastricht, The Netherlands
4 Department of Plastic and Reconstructive Surgery, Erasmus University Medical School, PO Box 1738, 3000 DR Rotterdam, The Netherlands
*Author for correspondence (e-mail: vandeneijnde{at}molcelb.unimaas.nl)
Accepted July 5, 2001
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SUMMARY |
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Key words: Myotube formation, Skeletal muscle development, Heart development, Apoptosis, Mouse embryo
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INTRODUCTION |
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At present, the molecular control of myogenesis has been studied in most detail in Drosophila, in particular for skeletal muscle (Dobberstein et al., 1997). In this species the genes myoblast city, blown fuse, rolling stone and Drac1G12V have been shown to be essential to myotube formation. These genes encode proteins that mediate key processes of recognition and adhesion and formation of a prefusion complex, as well as plaque, cell alignment and plasma membrane apposition and plasma membrane breakdown, respectively. Molecules that have been implicated in mammalian skeletal muscle differentiation include active protease nexin, Ca2+, cathespin B, desmin, GRP49, ERK6, m-calpain, NCAM, N-cadherin, proteasomes and the H145 antigen (Crescenzi et al., 1994; Dourdin et al., 1999; Dourdin et al., 1997; Gogos et al., 1996; Gorza and Vitadello, 2000; Hyodo and Kim, 1994; Lechner et al., 1996; Li et al., 1994; Moncman and Wang, 1998; Peck and Walsh, 1993; Seigneurin-Venin et al., 1996). Extending our knowledge of intercellular interactions in vertebrate muscle development may aid in the understanding of muscle tissue repair, which includes the reassembling of intercalated disks in the infarcted or hibernating heart (Kaprielian et al., 1998; Matsushita et al., 1999), and the fusion of satellite cells with damaged myotubes in skeletal muscle after exercise (Anderson, 1998).
The aim of the present study was to explore in greater detail the role of cell surface exposure of PS in myoblast differentiation, both in mouse embryos in vivo, and in established muscle cell lines C2C12 and H9C2 in vitro. Because PS exposure is predominantly considered a hallmark of apoptosis, occurring downstream of changes in the mitochondrion and after caspase activation (Martin et al., 1995; Verhoven et al., 1999), we compared maturation-induced and apoptosis-associated PS exposure. To achieve this, the spatiotemporal pattern of annexin V binding was determined during myoblast differentiation in relation to a panel of differentiation and cell death markers. In addition, the function of surface-exposed PS in myotube formation was studied by fusion-inhibition studies using annexin V.
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MATERIALS AND METHODS |
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In vitro studies
Cell lines
Two muscle cell lines were used in this study, both obtained from the American Type Culture Collection (Manassas, VA): the mouse C2C12 skeletal muscle cell line and the rat H9C2(2-1) cardiomyocyte cell line (Su et al., 1999). Importantly, the latter cell line has been described to possess features of skeletal muscle differentiation, including myotube formation (Menard et al., 1999). This is in contrast to heart muscle cells in vivo that do not fuse into myotubes but become connected by intercalated disks. The cells were grown in a humidified incubator at 5% CO2 and 37°C in growth medium (GM) consisting of DMEM (ICN Biomedicals BV, Zoetermeer, The Netherlands) supplemented with 2 mM L-glutamine (Serva, Heidelberg, Germany), 10% FCS (Gibco, Paisly, UK) and 0.05 mg/ml gentamycin (AUV, Cuijk, The Netherlands). At 70-80% confluency, cells were trypsinized (0.125% trypsin (Gibco), 0.02% EDTA and 0.02% glucose in PBS) for 1-3 minutes and split at a 1:5-1:10 ratio. Myotube formation was induced by replacing GM with differentiation medium (DM) (Van der Loop et al., 1996). The only difference between GM and DM is that the latter contains 2% normal horse serum (Gibco) instead of 10% FCS. To limit autofluorescence, all the experiments were performed with cells maintained in GM or DM deficient in neutral red (ICN Biomedicals BV).
As a positive control for myoblast differentiation-dependent annexin V binding, the BHK-21/C13 cell line (Flow Laboratories, Irvine, UK) was used, which has been reported to exhibit myoblast like characteristics, including the formation of multinucleated myotubes (Van der Loop et al., 1996). As negative controls, the myeloid cell line U937 (American Type Culture Collection) and the non-small-cell lung cancer cell line MR65 (Gropp, Philips Universitäts Klinik, Marburg, Germany) were used.
Reagents
In this study, several variants of human recombinant annexin V were used: (1) human recombinant annexin V conjugated to Oregon Green (annexin V-fluo) at a final concentration of 250 ng/ml (annexin V-Oregon Green, NeXins Research BV); (2) unlabeled recombinant human annexin V (AnxV; 1-100 µg/ml); (3) its null mutant (M1234; 100 µg/ml), which has mutations in all four Ca2+ binding sites resulting in a loss of PS binding capacity (Mira et al., 1997); and (4) M1234 conjugated to Oregon Green (1 µg/ml).
To test cell viability and apoptosis, the following reagents were used: propidium iodide (PI, 5 µg/ml; Molecular Probes, Eugene, OR), CMXRos (Mitotracker® Red, 500 ng/ml; Molecular Probes); Hoechst 33258 (10 µg/ml; Molecular Probes), and zVAD(OMe)-fmk (100 µM, diluted in DMSO; Alexis Biochemicals, Leiden, The Netherlands).
For immunofluorescence studies, rabbit-derived antibodies were used against active caspase 3 (polyclonal antibody CM1, 1:40, Idun Pharmaceuticals Inc., La Jolla, CA), and against annexin V (1:100). Furthermore, mouse derived mAbs were used directed against the sarcomeric protein titin (9D10, 1:10; Developmental Studies Hybridoma Bank). As secondary antibodies, Texas-Red-conjugated swine anti-rabbit Ig, or rabbit anti-mouse Ig were used as appropriate (DAKO, A/S, Glostrup, DK). As negative controls, the primary antibody was omitted; all negative control samples showed an absence of immunoreactivity.
Immunocytochemistry
Cells cultured in the presence of annexin V-fluo were rinsed with ice cold annexin V binding buffer and thereafter fixed with 4% paraformaldehyde in annexin V binding buffer at 4°C, pH 7.4 for 10 minutes. Then the cells were rinsed twice with PBS, permeabilized for 10 minutes with 0.005% SDS in PBS at room temperature, rinsed with PBS containing 1% BSA and incubated at 4°C with CM1 antibody overnight, or one of the other antibodies for 2 hours. Subsequently, samples were rinsed with the same buffer and incubated for 2 hours with the appropriate fluorochrome-conjugated secondary antibody. After incubation with the secondary antibody, the samples were rinsed again and mounted with glycerol containing DAPI (Sigma Chemicals, St Louis, MO).
Annexin V binding assays
To test for cell surface exposure of PS, annexin V-fluo was added to the medium. The cells were maintained in culture for a period ranging from 15 minutes up to several days in a humidified 5% CO2 incubator at 37°C. For the longer culture periods, medium including annexin V-fluo was renewed every 2 days. Cells were studied upon binding of annexin V-fluo using an inverted fluorescence microscope with appropriate excitation and emission filters (Zeiss, Oberkochen, Germany; Leica Microsystems BV, Rijswijk, The Netherlands), Image acquisition was achieved using Ikaros (2.3) (MetaSystems, Heidelberg, Germany) or Openlab (Improvision, Lexington, MA) software. Images and composite figures were prepared using Adobe PhotoShop (5.0.2) and Illustratator (8.0) software, respectively (Adobe Systems Inc., San Jose, CA). To some images, deconvolution software was applied to remove out-of-focus information using the Openlab package (Improvision).
Myotube fusion-inhibition assay
To test whether recombinant human annexin V could inhibit myotube formation, C2C12 and H9C2 cells were grown in 96 well plates (µClearTM black tissue culture microplates, Greiner Labortechnik, Frickenhausen, Germany). When the C2C12 cultures had reached 25% or 50% confluency and the H9C2 cultures had reached 95% confluency, myoblast differentiation and myotube formation was induced by replacing GM with DM containing AnxV (1-100 µg/ml) immediately after the medium switch. Medium with the same amount of annexin V was refreshed twice a day for the lower doses (1-40 µg/ml) and every third day for the highest dose (100 µg/ml). To ascertain that annexin V interacted with the myoblasts in a PS-dependent manner, we used the non-PS-binding annexin V mutant M1234 at a concentration of 100 µg/ml. As a positive control, cells were incubated with DM without AnxV or M1234.
On DMd0 and at the end of the culture period (i.e. DMd5 for C2C12 and DMd11 for H9C2), dual interference contrast-microscopy images were captured of each well. After culturing, cells were fixed in 4% paraformaldehyde in annexin V binding buffer and stained for the sarcomeric protein titin as described above. Subsequently, the complete bottom of the wells with cells attached were cut out with a scalpel and mounted on a coverslide using glycerol/DAPI. In each sample, all multinuclear cells were identified by titin staining and their nuclei were counted on a Zeiss microscope using the 40x 1.2 NA oil objective. For each well, the total number of multinucleated cells and nuclei therein, and the ratio between both were calculated. Averages±s.e.m. were calculated for the data in each group (control, M1234 and AnxV), using MS Excel 98 (Microsoft, Redmond, WA). To test for significance, the Mann-Whitney (non-parametric) test was applied using SPSS (10.07a) for Macintosh (MacKiev, Cupertino, CA).
Control experiments
To determine whether fluorescence observed in differentiating H9C2 and C2C12 myoblasts was due to the presence of annexin V-fluo and not of unconjugated fluorochrome or autofluorescence, annexin V was immunocytochemically visualized. Using the same immunocytochemical procedure, it was verified whether unlabeled human recombinant annexin V had bound to differentiating myoblasts and the level of endogenous annexin V expression in C2C12 and H9C2 cells was assessed. Expression of annexin V was not detected in C2C12 cells, whereas, in apoptotic H9C2 cells, some endogenous annexin V expression was observed. To quantify the amount of annexin V in this cell line, an ELISA was performed, according to the manufacturers instructions (Zymutest, Hyphen Biomed, Andressy, France). Cell lysates were obtained by removing the medium, adding 100 µl of lysis buffer (10 mM Tris, 1 mM EDTA, pH 7.4) and collecting the cells with a rubber policeman. Subsequently, the suspension was subjected to three freeze-thaw cycles, and stored at 70°C until simultaneous analysis. The levels of endogenous annexin V-protein measured in this cell line were, however, very low, with values ranging from 0.0028-0.0008% per total protein content.
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RESULTS |
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PS exposure in differentiating myoblast cultures
C2C12 and H9C2 muscle cell lines undergo differentiation after serum deprivation, as indicated by the process of cellular elongation, fusion into di- and trinuclear elongated cells and formation of extremely elongated multinucleated myotubes.
These C2C12 and H9C2 cells were able to bind annexin V-fluo transiently (Fig. 2). C2C12 muscle cells were found to bind annexin V within 8 hours after switching from GM to DM (Fig. 2A1,A2). After 2 days, binding was maximal with approximately 60% of the cells positive for annexin V. By contrast, proliferating C2C12 muscle cells (Fig. 2A3) and myotubes (Fig. 2A4) after 8 days in DM were not labeled with annexin V-fluo. H9C2 cells behave similarly (Fig. 2B1-4), except that the first binding of annexin V-fluo was observed after 2.5 days in DM (Fig. 2B1,B2), was maximal after 8 days (on average 40% of the cells) and absent again after 12 days. Mitotic cells (Fig. 2B3) and myotubes (Fig. 2B4) were negative for annexin V-fluo. In annexin V-positive cells the distribution pattern of this marker changed time dependently. Between 15 minutes and 2 hours of incubation the annexin V-fluo labeling was seen at the cell surface, in-between cells. After longer incubation periods the annexin V-fluo became gradually internalized, as could be demonstrated by rinsing with Ca2+-depleted medium (which dissociates annexin V from cell surface-exposed PS), resulting in only a partial loss of the annexin-fluo labeling.
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Since myoblast differentiation is a highly organized process, we wondered whether the temporal window of annexin V labeling is related to a particular phase of myoblast differentiation, as for example indicated by the molecular organization at the sarcomeric level. To this end, cells were double-stained for annexin V and titin, which has been shown to be one of the earliest proteins to become expressed and organized in the developing sarcomere (van der Ven et al., 1993). Essentially, the labeling patterns in C2C12 and H9C2 cells were similar (Fig. 3). Double labeling for both markers was mainly observed at a phase when titin was expressed in dot-like aggregates (Fig. 3A1,B1). By contrast, virtually all cells exhibiting a filamentous titin organization were negative for annexin V-fluo (Fig. 3A2,B2).
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Annexin V fusion-inhibition studies
To investigate a possible causal relationship between PS exposure and myoblast fusion, we analyzed whether annexin V can inhibit the formation of myotubes in differentiating C2C12 and H9C2 cells. C2C12 cells grown to 25% or 50% confluency (Fig. 5A1,A2) and H9C2 cells grown to 95% confluency (Fig. 5B1,B2) were induced to differentiate by medium switch. We also induced C2C12 myoblast differentiation in cells grown to 80-100% confluency. However, in these cultures the numbers of myotubes that had formed at DMd5 were too high to permit accurate counting of myotube numbers and nuclei.
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For the quantitative and statistical analysis of differences in myotube formation between myoblasts cultured with or without AnxV, we counted all myotubes and nuclei therein in 5-17 wells per group. To this end C2C12 cells at DMd5 (Fig. 5A5) and H9C2 cells at DMd11 (Fig. 5B5) were labeled for titin to identify differentiated multinucleated cells, and DAPI to count the nuclei. Fig. 6A illustrates the differences in myotube numbers containing a given number of nuclei for C2C12 cells that were induced to differentiate when grown to 50% confluency and H9C2 cells grown to 95% confluency before switching to DM. It is evident from this figure that for C2C12, AnxV significantly reduces the number of myotubes compared with control and M1234 incubations, whereas for H9C2 this inhibitory effect is less pronounced
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Generally these observations strongly indicate that recombinant human annexin V inhibits myotube formation by homotypic interaction between muscle cells via surface-exposed PS.
PS-exposing myoblasts: differentiation or apoptosis?
PS exposure in apoptotic cells is suggested to be a downstream effect of mitochondrial changes and activation of the caspase cascade in the apoptosis signaling pathway. To establish whether a comparable molecular mechanism underlies PS exposure in fusing myoblasts, these events were investigated during in vitro myoblast differentiation (Fig. 7).
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DISCUSSION |
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Muscle cells transiently expose PS during differentiation
In this study we present data showing that, during embryogenesis, morphologically viable myoblasts transiently bind annexin V, and thus express PS at particular stages of development. These cells were, however, not in contact with phagocytes, nor were any phagocytes detected in their vicinity, as would be expected when these cells were apoptotic (Abood and Jones, 1991; van den Hoff et al., 2000). In the underlying study, we have focussed on skeletal muscle development. In this tissue, binding of annexin V appeared to be a synchronous process that was present at E13 without the accumulation of pyknotic cells. Annexin V was not observed anymore at E14 or later, when most primary myotubes have formed, and was preceding the phase of massive death of primary myotubes that occurs after E15 (Ashby et al., 1993a).
Since, in the organism, cell surface exposure of PS is a trigger for phagocytic removal of apoptotic cells (Fadok et al., 1992b; van den Eijnde et al., 1999), one may wonder how PS-exposing, differentiating myoblasts remain unnoticed by phagocytes. Macrophages show tissue specificity with respect to the receptors used for clearance of apoptotic cells. In E13 mouse embryos, shielding of PS specifically on apoptotic neurons using annexin V was found to inhibit phagocytosis, whereas apoptotic mesenchymal cells in the limbs were ingested by phagocytes (van den Eijnde et al., 1999). This finding corroborates studies by Fadok and co-workers, who showed that depending on their activation status, macrophages may recognize apoptotic cells in a PS-dependent or -independent manner via integrins (Fadok et al., 1992a). Another mechanism may relate to the observations that apoptotic cells can attract phagocytes actively by secreting chemotactic factors (Knies et al., 1998). PS-exposing viable muscle cells may, by lacking such factors, avoid the attention of scavenging cells despite the fact that they bear a putative signal for cell removal (Knies et al., 1998; Wilson et al., 1996).
Similar to the in vivo situation, two populations of annexin V binding cells could be discriminated in C2C12 and H9C2 cell cultures, that is, apoptotic and differentiating muscle cells. In contrast to apoptotic cells, differentiating cells showed intracellular labeling when incubated with annexin V-fluo for longer time periods. Endocytosis of annexin V-fluo through pinocytosis can largely be excluded since co-labeling of annexin V-fluo with DiI (labeling of the membrane of pinocytic vesicles) showed hardly any overlap (S.M.v.d.E. et al., unpublished). The nature of the annexin V-fluo vesicles in differentiating muscle cells is not yet known. However, based upon their cellular localization they resemble: (1) the cytoplasmic vesicles found in the gap junction-like prefusion complex that forms at an early stage of myotube formation in Drosophila, directly following the initial phase of cell recognition and adhesion; and (2) the plasma membrane vesicles found during the phase of plasma membrane breakdown, which is believed to effect cytoplasmic continuity and actual fusion of myoblasts by removing excess plasma membrane in-between the fusing cells (Doberstein et al., 1997; Paululat et al., 1999).
In our in vitro experiments, annexin V was found to almost exclusively label mononucleated cells in contact with other mononucleated cells or small myotubes containing a few nuclei. Large, multinucleated myotubes were unlabeled. Labeling for the sarcomeric protein titin revealed further detail about the time-point of PS exposure in myoblast differentiation. Titin is the first sarcomeric protein expressed in muscle cells (Van der Loop et al., 1996; van der Ven et al., 1993); in differentiating muscle cells titin organization changes from diffuse to punctate, thereafter becomes filamentous and finally incorporates into sarcomeres, where it exhibits the typically striated staining pattern. Annexin V-positive myoblasts were largely characterized by a punctate titin distribution pattern, while subsequent stages of titin patterns were rarely observed in these labeled cells. In view of previous immunocytochemical studies of C2C12 and H9C2 cells (Van der Loop et al., 1996) or in human skeletal muscle cell cultures and BHK cells (van der Ven and Furst, 1998; van der Ven et al., 1993), our results indicate that, in particular, early postmitotic myoblasts expose PS. Thus, these in vitro data corroborate our in vivo observations that annexin V binding is mainly restricted to myoblasts in the process of myotube formation, and that PS is therefore transiently exposed and internalized before the fusion process is actually completed. The localization of PS in the inner plasma membrane leaflet in most cells is maintained by an aminophospholipid translocase (Diaz and Schroit, 1996; Zwaal and Schroit, 1997). During apoptosis and platelet activation, PS becomes cell surface exposed by a simultaneous translocase inhibition and scramblase activation (Higgins, 1994; Verhoven et al., 1995). It remains to be resolved, however, whether the same or similar transporters regulate loss of PS asymmetry of the plasma membrane in developing myotubes.
PS exposure during myogenesis is not related to the molecular cascade of apoptosis
A possible link between the apoptotic pathway and PS exposure during muscle cell differentiation was investigated and could be excluded at the level of loss of mitochondrial inner cell membrane potential, caspase 3 activation and caspase activity. In contrast to apoptotic H9C2 and C2C12 cells, annexin V-positive differentiating cells showed none of these molecular characteristics. In addition, we were unable to inhibit myotube formation and differentiation-dependent PS exposure with the broad spectrum caspase inhibitor zVAD(OMe)-fmk, suggesting that caspases are probably not directly involved in the loss of plasma membrane PS asymmetry. Nonetheless, we cannot rule out that PS exposure by myoblasts is triggered by a short and transient loss of mitochondrial membrane potential (Minamikawa et al., 1999). We consider it likely that PS exposure during muscle development is regulated via factors upstream of the mitochondrion. This hypothesis is substantiated by several recent studies, indicating that PS exposure does not always require caspase activity, and that this plasma membrane alteration can be reversed (Hammill et al., 1999; Williamson et al., 1995).
Physiological function of PS exposure in myoblast interactions
Using the merocyanin 540 dye for labeling of loose lipid packing of plasma membranes, Sessions and Horwitz, as early as 1981, suggested that in primary cultures of chick and quail myoblasts PS was exposed at the cell surface when these were expected to fuse (Sessions and Horwitz, 1981). From these results, they hypothesized that an increase in the percentage of the PS in the outer plasma membrane leaflet may give rise to an increase in membrane fluidity. In turn, this increased membrane fluidity could create a biochemical microenvironment facilitating the fusion process. Data presented in this study also indicate that cell surface-exposed PS is required for differentiating muscle cells to fuse into multinucleated myotubes. A significant decrease was observed in the numbers of myotubes formed in myoblast cultures containing 40 and 100 µg/ml of recombinant human annexin V. Generally, C2C12 cells showed more prominent features of differentiation than H9C2 cells. This may relate to the fact that C2C12 cells are of skeletal myoblast origin, whereas H9C2 cells derive from cardiac myoblasts. Cells from this lineage normally become connected via intercalated disks and do not form myotubes, neither in vivo nor in primary cultures (Kostin et al., 1999). It may be expected, therefore, that H9C2 cells are at least partially deficient in the expression of molecules regulating myoblast fusion and are less efficient in executing this process. In all experiments, a limited effect of the annexin V mutant M1234 on myotube formation was observed, which was borderline-significant in H9C2 cell cultures. This finding suggests that this annexin V mutant, which does not bind PS that is present in artificial bilayers of phosphatidylcholine (C.P.M.R., unpublished), may have a residual interaction with PS in biological membranes. Alternatively, it may suggest a PS-independent interaction of M1234 with myoblasts. At present, the nature of these interactions remains to be resolved.
Data from the fusion inhibition experiments indicate that PS is of physiological importance to myotube formation. A well documented function of PS is phagocyte recognition of apoptotic cells (Fadok et al., 2000; Hamon et al., 2000; Schroit et al., 1985; Verhoven et al., 1995), involving PS exposure at the plasma membrane of both scavenger and prey (Marguet et al., 1999). Cell-cell recognition is considered the initial event in myotube formation (Doberstein et al., 1997; Paululat et al., 1999), and we therefore consider it likely that PS has an analogous function in phagocytosis and myotube formation. In this respect, the phenomenon of partial inhibition of myotube formation by high doses of annexin V may relate to the timing of interference with the fusion process, at which a fraction of the cells may have already completed their recognition phase. Interestingly, a similar partial inhibitory effect of has been observed in studies on the role of PS in phagocytic removal of apoptotic cells. In several phagocyte lineages the decrease in phagocytic activity also maximized at approximately 50%, irrespective of the inhibitory agent used [i.e. annexin V at a dose 18-180 µg/ml (Bennett et al., 1995), PS-containing liposomes, or antibodies against a putative PS receptor (Fadok et al., 2000)]. This partial inhibition of phagocytosis is explained by the existence of multiple pathways in which PS is a membrane component amongst others orchestrating the engulfment.
A function of PS in homotypic cell-cell interactions would fit our observations that PS is initially exposed at the plasma membrane at myoblast cell-cell contact areas by early differentiating muscle cells. In addition, it could explain our observation of a transient exposure of PS in the developing heart in mouse embryos at E12 (Fig. 8). In heart development, homotypic cell recognition is possibly of importance to the formation of intercalated disks. PS exposure was detected, in particular, in sub-epicardial ventricular cardiomyocytes and the smooth-walled atrial myocardium in a spatiotemporal distribution, which did not match early descriptions of cell death foci in the developing heart (van den Hoff et al., 2000).
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Taken together, our studies in combination with those of others strongly indicate a physiological role of cell surface exposure of PS in myoblast differentiation.
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ACKNOWLEDGMENTS |
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