Department of Cell Biology and Anatomy, Cardiovascular Developmental Biology Center, Medical University of South Carolina, 173 Ashley Avenue, Charleston, SC 29425, USA
*Author for correspondence (e-mail: kubalaks{at}musc.edu)
Accepted 1 November 2001
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SUMMARY |
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Key words: Apoptosis, Outflow tract, Retinoid X receptor, TGFß2, Whole mouse culture
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INTRODUCTION |
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Proper growth and maturation of the heart requires a variety of molecular signals including retinoic acid (RA) (Chien et al., 1993; Fishman and Chien, 1997
; Kubalak and Sucov, 1999
). Too much or too little RA causes a spectrum of cardiovascular malformations including OFT and aortic arch anomalies (Lammer et al., 1985
; Pexieder et al., 1995
; Taylor et al., 1980
; Wilson and Warkany, 1949
; Wilson et al., 1953
), AV canal malformations (Wilson and Warkany, 1949
; Wilson et al., 1953
), looping disturbances (Dickman and Smith, 1996
; Smith et al., 1997
) and ventricular septal defects (Smith et al., 1998
; Wilson and Warkany, 1949
; Wilson et al., 1953
). Still, the mechanistic links between the teratogenic effect of RA and the resultant anomalies remain clouded by the fact that RA activates multiple signaling pathways.
A variety of gene-targeted mice have been generated to investigate the roles of RA during development. Mutations in several of the RA receptor subtypes lead to specific cardiac malformations and provide an avenue for understanding both the events leading up to embryonic lethality and for investigating mechanistic links between RA signaling and cardiac malformations (Kastner et al., 1994; Kastner et al., 1997
; Sucov et al., 1994
). Previous studies have documented the wide range of OFT abnormalities in the embryonic day (E) 13.5-15.5 retinoid X receptor
knockout (Rxra/) mouse (Gruber et al., 1996
). The significance of RXR
in heart development is underscored by the fact that embryos heterozygous for Rxra also displayed cardiac malformations (Gruber et al., 1996
). Defects were observed in several compartments of the heart, including the aortic sac, conotruncus, atrioventricular canal and ventricular myocardium (Gruber et al., 1996
). To date, no candidate downstream genes have been uncovered that offer an explanation of how these cardiac malformations occur.
We report that maldevelopment of the aorticopulmonary (AoP) septum is evident by E11.5 in the Rxra/. By E12.5, malformations of the endocardial cushions in the OFT became apparent. Associated with the malformed cushions is an increased level of apoptosis. In normal development, the onset of apoptosis in OFT cushion tissue occurs between E11.5 and E12.5 (for a review, see (Poelmann et al., 2000). Malformations in this region were evident by E12.5, coincident with the apoptosis and at least 1 day before the embryonic lethality in mutant mice. Concomitant with these morphologic changes was elevated levels of transforming growth factor ß2 (TGFß2) protein in the Rxra/ embryonic heart. When we exposed wild-type whole mouse embryos in culture to TGFß2 protein, apoptosis was enhanced in the OFT. TGFß2 treatment also lead to abnormal development of the OFT cushions and AoP septum. Accordingly, lowering TGFß2 levels by intercrossing the TGFß2 heterozygous mouse into the Rxra/ background partially restored the apoptosis and decreased the developmental defects. These results suggest that elevated levels of TGFß2 can increase apoptosis in the OFT, which in turn perturb cardiac development. This study is the first report that identifies a potential downstream target molecule in the retinoic acid signaling pathway proven to be critical during cardiac morphogenesis.
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MATERIALS AND METHODS |
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Gene expression analyses
RNase protection analysis and semi-quantitative reverse transcriptase-polymerase chain reaction (RT-PCR) were performed on single hearts that had only their atria removed (Gruber et al., 1996; Kubalak et al., 1994
). Preparation of RNA and generation of the MLC2a probe was as previously described (Kubalak et al., 1994
). The TGFß1 and TGFß2 plasmids were kind gifts from Harold Moses and probes were generated as described (Derynck et al., 1986
; Miller et al., 1989
). For RT-PCR, RNA was prepared as described previously (Kubalak et al., 1994
). PCR amplification of a 132 bp fragment corresponding to wild-type TGFß2 was as described by Sanford et al. (Sanford et al., 1997
). Amplification conditions were optimized to acquire semi-quantitative expression levels of TGFß2 and GAPDH (as control). The conditions for TGFß2 were 30 cycles at 95°C for 30 seconds, 50°C for 50 seconds and 72°C for 1 minute. PCR primers for GAPDH were as described by Kondo et al. (Kondo et al., 1998
) and the conditions were 20 cycles at 94°C for 30 seconds, 54°C for 30 seconds and 72°C for 45 seconds. Relative expression levels of TGFß2 were normalized to GAPDH using densitometry (NIH Image 1.62) of the respective PCR products. Numbers represent the -fold expression relative to wild type.
Whole-mouse-embryo culture
Preparation of rat serum was performed as described (Cockroft, 1990). Whole mouse embryos were cultured as described previously (Gruber et al., 1998
; Hertig et al., 1999
). Embryos were incubated in culture for 1 hour before the addition of reagents and then treated either with vehicle [4 mM HCl, 0.1% bovine serum albumin (BSA)] or with TGFß2 protein (R&D Systems, Minneapolis, MN) at a final concentration of 0.1, 10 or 30 ng/ml. The culture continued for 18 hours at which time the media was changed to fresh, reagent-containing, pre-equilibrated media and allowed to continue for an additional 6 hours. The embryos were then fixed in 2% paraformaldehyde-PBS for 1 hour at room temperature, washed in PBS and embedded as described above.
Immunohistochemistry
Protein expression was examined by first rehydrating the sections through xylenes and graded ethanols, then rinsing in PBS. Sections were blocked in 10% normal goat serum for 1 hour, washed in PBS, and incubated with antibodies against either TGFß1 (1:100), TGFß2 (1:300) or TGFß3 (1:500) (Santa Cruz Biotechnology, Santa Cruz, CA). They were then washed in PBS and incubated with FITC-conjugated goat anti-rabbit secondary antibody for 2.5 hours at room temperature. All primary and secondary antibodies were diluted in 0.1% BSA-PBS. After a final wash in PBS, slides were mounted with Slowfade AntiFade (Molecular Probes, Eugene, OR) and analyzed using a confocal laser-scanning microscope (BioRad MRC-1024, Cambridge, MA).
BrdU incorporation
Timed pregnant mice at E12.5-14.5 were injected intraperitoneally with 100 µg/gm body weight of 5-bromo-2'-deoxyuridine (BrdU, Sigma) in 200 µl of sterile 0.9% NaCl 1 hour before sacrificing (Porteus et al., 1994). Embryos were dissected from the uterus and processed as described above for paraffin-wax embedding and sectioning in the transverse orientation. After rehydrated through graded ethanols, sections were washed in TBSA-BSAT (10 mM Tris-HCl, 150 mM NaCl, 0.02% sodium azide, 1% BSA, 0.1% Triton X-100). They were then incubated with 4 N HCl for 30 minutes, followed by neutralization with 100 mM sodium tetraborate for 5 minutes, and then incubated in 0.1% Triton X-100 for 20 minutes. Specimens were co-stained with monoclonal anti-BrdU (Developmental Studies Hybridoma Bank, Iowa City, IA) and polyclonal atrial myosin light chain 2 (MLC2a, 1:500) (Kubalak et al., 1994
) in TBSA-BSAT, for 16 hours at room temperature. After a wash in TBSA-BSAT, they were incubated with FITC-conjugated anti-mouse secondary antibody and Cy5-conjugated anti-rabbit at room temperature for 2.5 hours. After a final rinse in TBSA-BSAT, the sections were mounted with Slowfade AntiFade mounting media and analyzed by laser-scanning confocal microscopy.
TUNEL staining
TUNEL assays were performed using the ApopTag kit (Oncor, Gaithersburg, MD) according to the manufacturers recommendations with minor modifications in order to co-stain for MLC2a. After incubation in the TdT enzyme reaction mixture, sections were exposed to fluorescein-conjugated anti-digoxigenin combined with anti-MLC2a for 30 minutes at room temperature. Slides were then washed for 15 minutes in PBS. After a 1 hour incubation with anti Cy5-conjugated anti-rabbit secondary antibody in 1% BSA-PBS, the slides were mounted as described above.
Quantitation of apoptotic cells in the sinistroventralconal cushion (SVCC) and the dextrodorsalconal cushion (DDCC) was estimated by scoring the number of apoptotic nuclei in serial sections 40 µm apart from the most anterior to the most posterior aspects of the cushions. Because in Rxra/ hearts it was frequently difficult to discern valve primordia from other neighboring cushion tissues, all sections in the series that contained cushion tissue were routinely included in the TUNEL analysis in both wild-type and mutant embryos. Apoptotic nuclei in the myocardium and endocardium of the OFT were excluded from the quantification. Positive nuclei were normalized to the total number of cushion cells counted from adjacent Hematoxylin and Eosin sections. Data are expressed as the number of positive apoptotic nuclei as a percentage of total cushion cells. Differences were considered significant when P<0.05 using a two-way analysis of variance.
Caspase activity assay
Caspase activity was performed using the Caspatag Fluorescein Caspase Activity Kit (Intergen Co., Purchase, NY) according to the manufacturers recommendations with minor modifications to assay single embryonic hearts. Briefly, cardiac cells from E12.5 embryonic hearts (atria removed) were isolated by incubation and trituration in 0.05% trypsin for 20 minutes at 37°C. Activated caspases were then labeled and quantitated by measuring the intensity of the fluorescein label using a fluorescence spectrophotometer (SLM Aminco, Urbana, IL). A portion of the final cell suspension was removed for protein determination in order to normalize the caspase activity to total protein using the Micro BCA Protein Assay Kit (Rockford, IL). Each litter was assayed separately and results are expressed as -fold change relative to wild-type embryos within each individual litter. Statistics were performed on -fold change values with differences considered significant when P<0.05 using a one-way analysis of variance.
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RESULTS |
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By E12.5 in the wild type, the opposing DDCC and SVCC ridges have made contact with each other and, in the majority of specimens studied have begun to fuse (Fig. 1I,K, arrow), indicating that septation of the OFT is well under way by E12.5. At this age in the Rxra/, the development of both OFT cushions was compromised. Not only was there a lack of fusion of the cushions (Fig. 1J,L), we also frequently observed evidence of abnormal interactions between the AoP septum and the conotruncal ridges, as was suggested at E11.5. Indeed, by E12.5 all of the Rxra/ embryos examined had malformed OFT cushions (11/11) and of those, 64% (7/11) displayed a lack of any fusion between the AoP septum and the SVCC and DDCC suggestive of an imminent AoP window (Fig. 1L, arrow). The remaining embryos displayed incomplete fusion resulting in irregular and hypoplastic ridges (Fig. 1J,L). To evaluate the relative sizes of the DDCC and SVCC in wild-type and mutant embryos, we quantitated the number of cushion cells in sections every 40 µm apart, spanning from the most distal to the most proximal aspects of each of the cushions (Fig. 1M). Confirming what we observed visually, the relative number of cushion cells in both the DDCC and SVCC was significantly less in the mutant (669±116 and 761±69, respectively) when compared with the wild type (1170±107 and 1113±105, respectively, P<0.01) (Fig. 1N). Therefore, by E12.5, the Rxra/ is characterized by an underdeveloped AoP septum and hypoplastic OFT cushions.
By E13.5 in the wild type, the DDCC and SVCC have nearly completely fused (Fig. 2A) and OFT morphogenesis proceeds into the final stages of septation. In the mutant, the two cushions are so significantly malformed and undersized that the lack of fusion may simply reflect they are physically too far apart to make proper contact with each other (Fig. 2D). The varying degrees of fusion and hypoplastic nature of the OFT ridges would explain the wide range and severity of the OFT defects typically seen in the Rxra/ (Gruber et al., 1996). Collectively, the data demonstrate that the morphological manifestations of a lack of RXR
protein in the OFT and aortic sac are evident between E11.5-12.5, at least 1 day before the embryonic lethality in these mice.
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Myocardialization is the process of myocyte migration that contributes to the formation of the muscular portion of the outlet segments (van den Hoff et al., 1999) and begins at E12.5-13.5 in the mouse (Waller et al., 2000
). These specialized myocytes have been reported to co-express MLC2a and MLC2v (Franco et al., 1999
). To determine if myocardialization is occurring in the Rxra/ mice, sections were selected from E13.5 embryos in order to visualize the area between the primordia of the aortic and pulmonary outlets. Identification of this region in wild-type embryos is straightforward as the two outlet cushions are well fused at this stage of development and the condensed mesenchyme is readily identified on histological sections (Fig. 3A, broken line). However, in the Rxra/, identification of the corresponding region is complicated by the frequent occurrence of unfused, hypoplastic conotruncal ridges (Fig. 3B). Therefore, serial sections 40 µm apart were evaluated to ensure the analysis of any potential regions of myocardialization. Interestingly, MLC2a and MLC2v were not co-expressed in all myocardializing myocytes in either wild-type or mutant embryos at E13.5. The lateral myocardial cuffs of the OFT expressed both MLC2a and MLC2v (Fig. 3C-F asterisks). However, the midline of this region in the wild type (Fig. 3C, arrowheads) and the corresponding region in the mutant (Fig. 3F, arrowheads) expresses only MLC2v. Importantly, myocytes along the endocardial cushion-myocardial interface manifest the finger-like phenotype characteristic of myocardializing myocytes. This was particularly evident in the wild type (Fig. 3G, arrows). In the Rxra/, even though the OFT cushions were not fused, a few myocytes could still be identified with the same finger-like phenotype projecting into the OFT cushions (Fig. 3H, arrows).
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At E13.5 in the wild-type heart, apoptosis is predominantly confined to OFT cushion tissue lying between the aortic and pulmonary outlet valve primordia where the two OFT cushions have fused (Fig. 3E) (Okamoto et al., 1981; Ya et al., 1998
). Atrioventricular cushion tissue and the myocardial crest of the interventricular septum also contain apoptotic nuclei which is normal at this stage of development (Poelmann et al., 2000
). The sites of cell death in the mutant heart were confined to these same regions (Fig. 3F, and data not shown). Thus, ectopic apoptosis was not found in the Rxra/ endocardium, epicardium or the myocardium of either the ventricles or atria (data not shown).
As E12.5 mutant embryos already display hypoplastic cushions (Fig. 1J,L) before any overt decrease in proliferation, we set out to evaluate whether this could be explained by a higher degree of apoptosis (Fig. 4). To quantitate this, we counted the number of apoptotic nuclei in serial sections 40 µm apart from the most cranial to the most caudal aspect of the OFT cushions (Fig. 1M). Fig. 4 shows representative images from a single wild-type embryo (Fig. 4A,C,E) and a single mutant (Fig. 4B,D,F). In wild-type embryos, 3.9% of cells in the DDCC and 5.5% of cells in the SVCC were apoptotic, while in mutant embryos this was elevated to 10.6% and 9.8%, respectively (Fig. 4G; P<0.05). Additionally, the region of the enhanced apoptosis in the mutant extended more cranial and caudal than in wild-type cushion, particularly in the SVCC (Fig. 4A-F). To confirm what we observed using the TUNEL assay, we analyzed caspase activity in trypsin-dispersed embryonic heart cells. Caspase activity has been reported to be a good indicator of apoptotic activity in the developing OFT (Watanabe et al., 1998). Consistent with the TUNEL assay, we observed significantly elevated caspase activity in mutant E12.5 hearts (Fig. 4H). Not surprisingly, as there were no apparent morphological differences between E11.5 wild-type and mutant OFT cushions, there were also no differences in the observed programmed cell death. In fact, there was essentially no OFT apoptosis for either genotype at E11.5 (data not shown). Hence, we conclude that elevated apoptosis is responsible for the hypoplastic phenotype of the OFT cushions observed in Rxra/ mouse.
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DISCUSSION |
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Effect of enhanced apoptosis on OFT development in the Rxra/
Each mutant E12.5 embryo examined displayed irregular and hypoplastic conotruncal ridges. We conclude that this is not a result of a decrease in proliferation as we could not detect differences in BrdU incorporation until E14.5 when the embryos were already showing the typical signs of lethality in these mice (Sucov et al., 1994). The hypoplastic phenotype could also be the result of a decrease in the number of cells that undergo epithelial-mesenchymal transformation. This, too, is unlikely as there were no obvious morphological differences between wild-type and mutant OFT regions through E11.5. Additionally, OFT cushion explants from E9.5 mutant embryos displayed equivalent epithelial-mesenchymal transformation and migration of mesenchymal cells as their wild-type littermates in three-dimensional collagen gel cultures (S. W. K., P. J. Gruber and K. R. Chien, unpublished). Indeed, at E11.5 in the Rxra/, the sizes of the cushions and relative density of cushion cells were similar to the wild type. Alternatively, enhanced apoptosis could lead to hypoplastic cushions by eliminating cells that would otherwise proliferate in maturing cushions. Neither wild-type nor Rxra/ embryonic cushions displayed any significant cell death at E11.5. However, at E12.5 apoptosis was significantly elevated in the Rxra/ over that in the wild type. Because, in normal development, apoptosis in the OFT progressively increases between E12.5 and E13.5, the enhanced apoptosis at E12.5 in the mutant may represent premature differentiation of cells predestined for programmed cell death. Chambon and co-workers have suggested this concept for the ventricular myocyte population in the Rxra/ embryo (Kastner et al., 1997
). The additional apoptotic cells in the OFT of the Rxra/ may be surrendering to this same fate. Alternatively, the enhanced apoptosis may represent a subpopulation of cells that would have otherwise not undergone programmed cell death. The exact identity of the apoptotic cell population is currently under investigation. Nonetheless, our results suggest that enhanced apoptosis, and not decreased proliferation, is the major contributing factor to the development of the irregular and hypoplastic cushions in the Rxra/.
The precise role of apoptosis during OFT development has not been established. Programmed cell death would not contribute to the overall structure of the OFT cushions per se, but instead may occur as a prerequisite to myocardialization (Poelmann et al., 2000). One line of thinking is that apoptosis of neural crest cells triggers the activation of latent TGFß2 and, in turn, signals adjacent OFT myocytes to begin myocardializing the apoptotic zone (Poelmann et al., 2000
). Conversely, our data support the notion that TGFß2 might be involved in a signaling cascade that would initiate, rather than respond to, the apoptosis. This does not rule out the possibility that cell lysis may activate latent TGFß2, particularly if a positive feedback loop exists between activated TGFß2 and programmed cell death. With regard to myocardialization, the fact that myocyte migration was not enhanced in either the Rxra/ or in TGFß2-treated whole mouse embryos suggests that TGFß2 may not be the stimulus that triggers myocyte migration during septation of the OFT. In fact, even though TGFß2 was elevated, myocardialization seemed to be blunted in the Rxra/. Interestingly, TGFß2-deficient mice also lack a muscular outlet septum (Sanford et al., 1997
). As fusion of the cushion ridges was also lacking in the Rxra/, it is intriguing to speculate that TGFß2, in combination with cushion fusion, may be required to trigger myocardialization. In either case, we propose that TGFß2 alone is not sufficient. Nevertheless, the temporal and spatial relationship between apoptosis and myocardialization leaves open the possibility that lysis of apoptotic cells somehow signals adjacent myocytes to begin their migration.
Identification of the cell type(s) that undergo apoptosis in wild-type embryos and enhanced apoptosis in the Rxra/ remains to be determined. The current line of thinking is that all cells destined to undergo programmed cell death in OFT cushion tissue are neural crest cells (Poelmann et al., 1998; Poelmann et al., 2000
). Our data do not discount this possibility, because the spatial distribution of apoptotic cells in the mutant is confined to the central region of the cushion ridges where the majority of neural crest cells are thought to reside. This would still be consistent with the sentiment that most dying cells in the mutant could be of neural crest origin rather than, for example, recently transformed epithelial cells (which would be found adjacent to the endocardium). However, the boundaries of apoptosis extend more cranial and caudal than in the wild type, suggesting that other cell types may be subject to programmed cell death in the Rxra/. It is intriguing to speculate that if lysis of neural crest cells is a prerequisite for myocardialization to occur (Poelmann et al., 2000
), then the fact that myocardialization is not enhanced in the Rxra/ may mean that the additional cells undergoing apoptosis in the mutant are not neural crest cells.
The developing OFT cushions also participate in the formation of the interventricular septum (Bartelings and Gittenberger-de Groot, 1991). At the inner curvature, the convergence of the outflow septum, interventricular septum, primary atrial septum, spina vestibuli and atrioventricular cushion tissue [collectively termed the central mesenchymal mass (Kirby and Waldo, 1995
)] culminates in the formation of a normally septated four-chambered heart (Kirby and Waldo, 1995
; Webb et al., 1998
; Wessels et al., 1996
). Therefore, it is not surprising that the Rxra/ has a high frequency (94%) of membranous ventricular septal defects (Gruber et al., 1996
) because apoptotic nuclei were found in the caudal aspect of the SVCC adjacent to the inner curvature. The lack of expansion of the cushion ridges in combination with the abnormal development of the ventricular myocardium would prevent the normal interactions between the outflow cushions and the interventricular septum and result in a membranous ventricular septal defect as seen in nearly every Rxra/ (Gruber et al., 1996
).
Contribution of TGFß2 to OFT and aortic sac development in the Rxra/
The transforming growth factor ß family of proteins has been shown to promote epithelial-mesenchymal cell transformation in culture. TGFß1 and TGFß3 can promote transformation of endothelial cells in cultured chick atrioventricular cushion explants (Potts and Runyan, 1989; Potts et al., 1991
). Data that support a role for TGFß2 in cushion development are limited, particularly in OFT cushion tissue. In the atrioventricular canal, it appears that TGFß2 and TGFß3 may substitute for each other in promoting epithelial-mesenchymal transformation (Potts and Runyan, 1989
). Perhaps the best data supporting a role for TGFß2 during OFT development have come from TGFß2 knockout mice, which display hyperplastic OFT cushions (Sanford et al., 1997
). This would imply that the lack of TGFß2 either decreases programmed cell death, or allows for a certain degree of cell division above normal. Surprisingly, however, it has been recently shown that TGFß2 knockout mice display an elevated level of apoptosis in the OFT, while proliferation was not determined (Bartram et al., 2001
). Embryos heterozygous for TGFß2 were apparently normal (Bartram et al., 2001
). Based on the phenotype of the TGFß2 mutant then, elevated TGFß2 should result in decreased cushion size. Our data are consistent with this concept because the OFT cushions were hypoplastic both in the E12.5 Rxra/ and in TGFß2-treated whole mouse embryos in culture. We propose that the effects of elevated TGFß2 on cushion development are due to enhancing apoptosis rather than decreasing proliferation.
Rxra/ embryos that were also heterozygous for Tgfb2 showed a partial restoration of the apoptosis and partial rescue of the developmental defects in the OFT. The reason for only a partial rescue may indicate that the precise level of activated TGFß2 in the Rxra//Tgfb2+/ varies between embryos. Alternatively, other molecules may also be involved, such that lowering TGFß2 alone may not be sufficient to rescue fully the defects in the Rxra/. Unfortunately, there is no reliable method to determine the precise levels of activated TGFß2 in an individual embryo. At present we are confident that TGFß2 plays a role in OFT remodeling; the extent of that contribution beyond modulating apoptosis is currently under investigation.
Normal development of the AoP septum results in the supravalvular formation of separate aortic and pulmonary outlets. If the AoP septum is either absent or incompletely formed, a spectrum of defects can be observed that ranges from persistent truncus arteriosus (no septum) to AoP window (incomplete septum). These defects are found in the Rxra/ (Gruber et al., 1996), as well as Splotch mice (Franz, 1989
) and neural crest ablation studies (Nishibatake et al., 1987
). However, very little information has been reported regarding the molecular regulation of the development of the AoP septum. Our data suggest that TGFß2 is involved in this process. Both Rxra/ embryos and TGFß2-treated whole mouse embryos displayed an abnormal development of the AoP septum. Importantly, in Rxra/ embryos that were also heterozygous for Tgfb2, the AoP septum had descended further down into the OFT and, in each embryo, demonstrated partial fusion with the OFT ridges. This supports the notion that TGFß2 contributes to the aortic sac defects in the Rxra/. The apparent partial rescue suggests that either there are other molecules working in concert with TGFß2, or the precise level of activated TGFß2 in the Rxra//Tgfb2+/ depicts the degree of rescue.
Taken together, the results suggest that disturbances in TGFß2 signaling disrupts the integrative processes of differentiation, apoptosis and proliferation within the OFT endocardial cushions and the aortic sac. Elevated TGFß2 results in enhanced apoptosis in OFT cushion tissue with minimal effects on proliferation until after the heart begins to fail. The increased apoptosis effectively removes cells from the cushions prematurely and results in hypoplastic cushions. As a result, the malformed DDCC and SVCC are prevented from achieving normal fusion and can not contribute to the septation of the OFT. The role TGFß2 plays in myocardialization may be indirect as myocyte migration appears to be decreased both in the Rxra/ where TGFß2 is elevated and in TGFß2-knockout mice where TGFß2 is absent (Sanford et al., 1997). Enhanced TGFß2 also attenuates the development of the AoP septum so that it can no longer interact properly with the OFT cushions. Consequently, defects that range from an AoP window to persistent truncus arteriosus would be observed depending on the degree of extension of the AoP septum toward the OFT.
We conclude therefore, that one role for Rxr during cardiogenesis is to maintain proper levels of TGFß2 expression in the OFT and aortic sac. TGFß2, in turn, modulates the degree of apoptosis during remodeling of the OFT cushions. Whether TGFß2 directly or indirectly promotes apoptosis of cushion cells remains to be determined. As TGFß2 receptors (type II) appear to be ubiquitously expressed in the heart and are unchanged in the Rxra/ (S. W. K., unpublished), the effects of elevated TGFß2 in the mutant may be more dependent on the levels of activated protein than on the expression of the receptors. Additionally, whether RXR
directly or indirectly regulates the TGFß2 expression during murine cardiogenesis is not known. In human osteoblasts and keratinocytes, RXR
heterodimerizes with vitamin D receptors to directly downregulate the TGFß2 promoter and inhibit cell growth in culture (Wu et al., 1999
). Therefore, it is possible that RXR
directly modulates TGFß2 expression in the embryo. However, an indirect regulatory mechanism is also plausible. Lack of RXR
in one cell may effect expression of TGFß2 in the neighboring cell. Alternatively, RXR
may indirectly regulate TGFß2 levels, owing to the potential contributions of TGFß2 protein by maternal sources or by the placenta. It has been shown in TGFß1 null mice that maternal sources of TGFß1 can rescue defects in the TGFß1 null embryo (Letterio et al., 1994
). Similar studies have not been carried out for TGFß2 null mice. Studies that have examined the maternal influence of TGFß2 on embryogenesis have primarily focused on the effects of TGFß2 on the developing fetal-maternal placenta (Ando et al., 1998
; Gorivodsky et al., 1999
; Schilling and Yeh, 2000
), rather than the maternal contribution of TGFß2 to the embryo. It has been shown that expression of TGFß2 in the fetal placenta protects the embryo from immune rejection by the mother (Gorivodsky et al., 1999
). Thus, a decrease in TGFß2 expression has been correlated with pregnancy loss (Gorivodsky et al., 1999
) while the effects of elevated expression have not been examined. Maternal TGFß1 has been reported to cross the placenta and can be found in the embryo (Letterio et al., 1994
). However, whether TGFß2 crosses the placenta and contributes to the development of the embryo is not known. Studies designed to evaluate the embryonic effects of a teratogenic insult on the placenta have thus far concentrated on the levels of TGFß2 mRNA and protein within embryonic target organs and not on potential circulating levels of growth factor from placental tissue (Ivnitsky et al., 2001
). Even assuming there are maternal contributions to the levels of TGFß2 exposed to the embryo, these contributions are probably minimal as we still observe differences in the levels of expression of TGFß2 protein between wild type and Rxra null mutants, even though both have a TGFß2 heterozygous mother. Additionally, mRNA levels were also elevated in the mutant heart. Our current work and that by Dickson et al. show that by E12.5 in the wild-type embryo, the levels of TGFß2 total protein have decreased in the heart to near undetectable levels, i.e. normal levels in the wild type (Dickson et al., 1993
). This suggests that altered levels of TGFß2 in the Rxra/ are largely determined by expression patterns within the embryo proper.
The role of RXR-related placental defects in the mutant must also be considered. Chambon and co-workers reported that the placenta of the Rxra/ was found to be abnormal from E14.5 (Sapin et al., 1997
). This suggests that placental defects may contribute only to the lethality from that day forwards. They also documented that premature differentiation of cardiac myocytes in the Rxra/, with regard to sarcomeric organization, is first observed at E9.5, before the overt placental defects (Kastner et al., 1997
). In our study, we observe malformations in the aortic sac as early as E11.5, also before overt placental defects. In a recent report describing the PPAR
null mouse, Barak et al. have described signs of abnormal development of the placenta in the E9.5 Rxra/ (Barak et al., 1999
). However, the authors also stated that the defects were not nearly as severe as those found in PPAR
null embryos. PPAR
null embryos die at E10.0 because of severe placenta-related developmental defects (Barak et al., 1999
). Indeed, no malformations other than the thin myocardium in the Rxra/ are manifest until E11.5 (our current study) or later (Dyson et al., 1995
; Gruber et al., 1996
; Sucov et al., 1994
). Because RXR
heterodimerizes with PPAR
(Kliewer et al., 1994
), one may expect to see an overlap in phenotype between these two knockout mice. However, Barak et al. have noted that the cardiac phenotype (thinning of the ventricular myocardium) in PPAR
null embryos markedly exceeds that observed in the Rxra null (Barak et al., 1999
) and, importantly, no other malformations in the heart were noted. This could indicate that the thin ventricular myocardium phenotype in the Rxra/ is due to inadequate PPAR
/RXR
signaling in the placenta, while the outflow tract and aortic sac phenotypes might occur via a distinct heterodimer pair such as RAR/RXR. Therefore, while sites of minimal placentation defects were observed at E9.5 (Barak et al., 1999
), overt defects that would effect the function of the placenta in the Rxra/ were not apparent until E14.5 (Sapin et al., 1997
). Moreover, it has been documented that the fetal-maternal circulatory system in the mouse does not become functional, with exchange of nutrients, until around E12.5 (Muntener and Hsu, 1977
). Collectively, these data suggest that at least the initial defects in the outflow tract and aortic sac of the Rxra/ occur independently of influences by the placenta.
Finally, identification of the cell types responding to the elevated TGFß2 remains to be established. Because during development of the murine ventricular myocardium, RXR functions in a non-cell autonomous manner (Chen et al., 1998
; Tran and Sucov, 1998
), it is possible that more than one cell type may be involved in the RXR
/TGFß2 signaling pathway that leads to the elevated OFT apoptosis. Candidate cell types include the neural crest cells, the mesenchymal cells of the cushions, as well as the endocardium, epicardium and myocardium. We are currently investigating which cell lineages are responding to the elevated TGFß2 in the Rxra/. Determining the regional locations where elevated TGFß2 becomes activated would help to answer this question, but the issue is complicated by the fact that presently available antibodies for immunohistochemistry can reliably detect only the latent form of the protein. A recent report in the chick heart holds promise for identifying regions where TGFß2 becomes activated (McCormick, 2001
). Identification of similar regions in the mouse will be important in order to predict what cell lineages may be under the influence of TGFß2.
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ACKNOWLEDGMENTS |
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