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Vol. 20, Issue 1, 256-269, January 1, 2009
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*Department of Cell Biology, Department of Biological Sciences, and Department of Medical Genetics, University of Alberta, Edmonton, AB T6G 2H7, Canada
Submitted March 17, 2008;
Revised October 14, 2008;
Accepted October 29, 2008
Monitoring Editor: Marianne Bronner-Fraser
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Cripps and Olson, 2002). Among these, myocyte enhancer factor-2 (MEF2) plays a key role in specification and subsequent differentiation of all muscle types (skeletal, smooth, and heart muscle; Lilly et al., 1995; Black and Olson, 1998). There are four different known vertebrate mef2 genes: mef2-a, -b, -c, and -d (Black and Olson, 1998). These four genes produce several different MEF2 isoforms involved in differentiation of all muscle types. In addition, it has been proposed that MEF2 proteins have a requirement for tissue-specific cofactors to confer additional specificity. For example, during mammalian heart development, GATA-4 (Charron and Nemer, 1999) helps to recruit MEF2 to the promoters of cardiac-specific genes including atrial natriuretic factor (ANF) and 
-cardiac actin (-CA; Morin et al., 2000). MEF2 also interacts with another transcription factor, HAND1, during activation of ANF in cardiac cells (Morin et al., 2005). This complex interplay between MEF2 proteins and cofactors is not restricted to cardiac muscles as during skeletal muscle development; MEF2 interacts with MyoD during activation of specific structural genes (Molkentin et al., 1995; Black et al., 1998).
In terms of MEF2 protein family activity, muscle differentiation in Drosophila is relatively less complex as there is only a single homologue mef2, Dmef2 (Lilly et al., 1994). Like vertebrates, Drosophila Dmef2 isoforms activate muscle-specific genes (Black et al., 1998; Black and Olson, 1998) and also seems to interact with a conserved cohort of interacting proteins for muscle specification, including cardiogenesis. These include tinman (Azpiazu and Frasch, 1993), dHAND (Han et al., 2006), and the gene encoding the GATA factor Pannier (Pnr; Gajewski et al., 1997). Dissection of the regulatory region of the muscle-specific structural genes, TroponinT (Butler and Ordahl, 1999), TroponinI (TnI), and Tropomyosin (TmI) indicates that cofactors work together with Dmef2 during cardiogenesis in Drosophila (Lin et al., 1996; Mas et al., 2004; Nongthomba et al., 2004). However, relatively little is known about the Dmef2 interacting partners during differentiation of somatic muscles (analogous to mammalian skeletal muscles) versus cardiac muscle cells. We have focused on the muscle-specific role of two proteins Scalloped (Sd) and Vestigial (Vg) that have been shown previously to be potent activators of fate specification in several nonmuscle cell types. There is considerable functional conservation in the activities of TEF-1/Sd and Vgl/Vg as mammalian TEF-1 can functionally substitute for Sd (Deshpande et al., 1997) and Vgl-2 can partially substitute for Vg during Drosophila development (Vaudin et al., 1999).
Sd is the only member of the Transcriptional Enhancer Factor-1 (TEF-1) family of proteins in Drosophila (Campbell et al., 1992) and together with an activating cofactor, Vg, induce formation of the wing. In fact, ectopic expression of Vg, in the cells of the developing eye that also express Sd, leads to a respecification of these cells to a wing phenotype (Halder et al., 1998; Simmonds et al., 1998). Vg has two domains that influence transcriptional activation activity (MacKay et al., 2003), and Vg requires Sd for nuclear localization (Halder et al., 1998; Simmonds et al., 1998; Srivastava et al., 2002). Both TEF-1 and Sd bind DNA via a conserved TEA domain, although like TEF-1, Sd does not exhibit significant transcriptional activation ability on its own. Vg interacts directly with Sd to form a transcription factor (TF) complex required for wing specific gene expression (Simmonds et al., 1998). There is also evidence that TEF-1 acts in concert with other transcription factors. For example, YAP65 has been identified as a powerful transcriptional coactivator of TEF-1 in mouse (Vassilev et al., 2001).
After identification of the Sd-interaction domain of Vg (Simmonds et al., 1998), several mammalian genes encoding Vestigial-like proteins with homologous domains were identified. These include Vestigial-like 2 (Maeda et al., 2002a), which interacts with TEF-1 in skeletal muscle to augment myosin heavy chain (MHC) expression (Maeda et al., 2002a; Gunther et al., 2004). Vestigial-like 4, which is enriched in heart muscle also functionally interacts with TEF-1 (Chen et al., 2004). Similarly, the Sd homologue, TEF-1 is an MEF2-interacting protein expressed in all muscle types (Stewart et al., 1994; Carlini et al., 2002). The phenotype of a TEF-1 mouse knockout suggests a role in cardiac maturation (Chen et al., 1994), but TEF-1 is also required for skeletal and smooth muscle gene expression (Pasquet et al., 2006). However, TEF-1 cannot activate transcription alone (Xiao et al., 1991), and overexpression of TEF-1 results in repression of transcription (Jiang and Eberhardt, 1996).
In terms of muscle development, mammalian TEF-1 has been shown to interact with MEF2, and this interaction interferes with MEF2-dependent activation of the β-Myocyte heavy chain (β-MHC) promoter (Maeda et al., 2002b). Other known MEF2 cofactors include poly-(ADP-ribose) polymerase (PARP) on the cardiac TnT gene (Butler and Ordahl, 1999), Max on the cardiac -myosin heavy-chain gene (Gupta et al., 1997), and serum response factor on the skeletal -actin gene (Gupta et al., 2001). Given the multiplicity of interactions between these proteins, it is possible that MEF2 and TEF-1 function within a larger complex of TFs that includes additional proteins, like members of the Vgl family and that alternative composition of these various complexes may provide cell-specific gene activation during muscle differentiation.
Although Dmef2 has a clear role in Drosophila muscle differentiation, specific functions for Vg or Sd in muscle cells has not yet been well characterized. To test the role for a complex of MEF2, TEF-1, and the Vgl-family of proteins in the differentiation of muscle cells led us to probe the combinatorial activities of each of these proteins during Drosophila embryonic muscle specification. There is some precedence for a role for Vg in muscle development as it had been reported to be required for late-stage development of indirect flight muscles (IFMs) derived from the wing disk–associated myoblasts (Sudarsan et al., 2001). In wing discs isolated from flies with null vg mutations, myoblasts proliferate, migrate, and fuse normally but further differentiation fails to occur (Bernard et al., 2003), a phenotype similar to that associated with mutations in Dmef2 (Nguyen and Xu, 1998; Nguyen et al., 2002). Although it is possible that this phenotype is due to the well known wing-specification role previously ascribed to Vg, it is equally possible that this represents a muscle-specific activity for Vg and Dmef2 and further suggests that these two proteins may functionally interact.
To clarify the role of Sd and Vg during embryonic muscle development, in addition to the IFM precursors, we have looked at all of the developing muscles in sd and vg Drosophila mutant embryos and found consistent defects in both the cardiac and somatic musculature. Additionally, we have shown that sd is expressed in at least some Drosophila embryonic muscles. Further, we have tested protein interactions between Drosophila Dmef2, Sd, and Vg and found that these proteins do interact both in vitro and in vivo. Finally, we have tested the specific combinatorial requirement for the presence or absence of Vg or Sd in certain muscle types because elevated expression of each causes significant defects in the specification or differentiation of specific muscle cell types.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
Drosophila Strains
Ectopic-expression of Gal4-UAS transgenes (Brand and Perrimon, 1993) was performed using Dmef2-Gal4 (Ranganayakulu et al., 1998), Sd-Gal4 (Roy et al., 1997), and 5053-Gal4 (Ritzenthaler et al., 2000). All other UAS-transgene animals were made in our laboratory for this study.
Plasmids
GST-Sd and GST-Dmef2 vectors were created by insertion of full-length sd and Dmef2 into the BamHI and SalI sites of pGEX-4T1 (GE Biotech, Piscataway, NJ), respectively. Vg deletions (see Figure 5) in pET16b (Novagen, Madison, WI) were as described previously (Simmonds et al., 1998). Expression vectors for transfection of S2 cells (see Figure 4) were created by Gateway Technology (Invitrogen) and the Drosophila Gateway destination vectors (Terrence Murphy, Carnegie Institute of Washington, Baltimore, MD).
Fluorescent In Situ Hybridization
Anti-sense digoxigenin (DIG, Roche, Indianapolis, IN) RNA probes targeting sd were made by creating a double-stranded PCR product with a T7 polymerase binding site incorporated into the 3' primer. The primers used were 5'-gaacaacctgagctgcagcgagttgg and 5'-taatacgactcactatagggagacagcacttggatgtgcg. Embryo fixation and hybridization of the probes and detection of the fluorescent signal were performed using the method of Hughes and Krause (1999), including the modifications outlined in Lecuyer et al. (2007).
Glutathione S-Transferase Pulldown Assays
Glutathione S-transferase (GST) fusion proteins were expressed in Escherichia coli [Rosetta 2(DE3), Novagen] and purified according to the manufacturer's directions (GE Biotech). Probe proteins were 35S labeled in vitro using the TNT-coupled in vitro transcription-translation system (Promega, Madison, WI). For the in vitro binding assay, 3–6 µl of 35S-labeled probe proteins were incubated with 2 µg of immobilized GST fusion proteins in 500 µl of buffer (20 mM Tris, pH 7.6, 100 mM NaCl, 0.5 mM EDTA, 10% glycerol, and 1% Tween-20) containing 0.25% bovine serum albumin (BSA) and protease inhibitor cocktail for 2 h at 4°C. The beads were washed six times in 500 µl of the same buffer, and the bound proteins were resolved by SdS-PAGE and analyzed by autoradiography.
Immunoprecipitations and Immunoblotting
S2 cells were transfected with relevant expression constructs containing the heat-shock promoter, and protein expression was induced by heat shocking cells for 35 min at 37°C. Cells were harvested 1 h after induction, washed one time in PBS, and resuspended in RIPA (radio-IP) buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 1.0% NP-40, 0.5% deoxycholic acid, 0.1% SdS, and protease inhibitor cocktail). The lysate was then incubated for 15 min at 4°C with agitation and centrifuged for 15 min at 13.2K rpm at 4°C, and the supernatant was then transferred to a fresh tube. CoIP reactions were carried out on 200 µl of supernatant (600 µl supernatant from a 25-cm2 flask of cell culture) using 8 µl anti-FLAG M2-agarose (Sigma, St. Louis, MO) in 500 µl RIPA buffer. Agarose beads were incubated for 1 h at 4°C with rocking, centrifuged for 1 min at 1.4K rpm at 4°C, and washed six times by vortexing in 500 µl RIPA buffer. Primary antibodies for immunoblotting were mouse anti-FLAG (1:1000; Sigma), rat anti-hemagglutinin (HA; 1:400; Roche), and rabbit anti-Myc (1:1000; Cell Signaling, Beverly, MA). Secondary antibodies were goat anti-mouse Alexa680 or IRdye800 (1:5000; Invitrogen), goat anti-rabbit Alexa 680 or IRdye800 (1:5000; Invitrogen), and goat anti-rat IRdye800 (1:5000; Invitrogen). Nitrocellulose membranes were scanned and analyzed by Odyssey Infrared Imaging System (Li-Cor Biosciences, Lincoln, NE).
Reverse Transcriptase PCR
Total RNA from stage 12–15 wild-type embryos and overexpression embryos was isolated with Trizol reagent (Invitrogen) and treated with DNase I (Ambion, Austin, TX). Reverse transcription was carried out using 2 µg of total RNA, SuperScript II reverse transcriptase (SS II, Invitrogen) and gene-specific first-strand primers. Subsequent amplification of the resulting cDNA was performed using Taq DNA polymerase (Invitrogen) and one pair of nested primers for each gene. Primers for the control rp49 cDNA were: first-strand primer, 5'-cttcttgagacgcaggcga and nested primers, 5'-agcatacaggcccaagatcg and 5'-agtaaacgcgggttctgcat. Primers for Act57B cDNA amplification were first-strand primer, 5'-gcaggagacaggtgagtagacc and nested primers, 5'-ctccggcatgtgcaagg and 5'-gcaacacgcagctcgttg. Primers for mhc cDNA amplification were 5'-agaaggctgaggaactgc and 5'-gttcaagttgcggatctg. The primers were designed for rp49 and mhc to span an intron and the forward primer for Act57B cDNA to span the conjunction of two exons. To make sure the RT-PCR was in the linear range of amplification, we performed PCR reactions at increasing cycle numbers (15, 20, 25, and 30), and similar results were observed. The RT-PCR was performed on two different mRNA isolations, and repeated three times, with consistent results.
Fluorescence Microscopy
Wild-type and overexpression embryos were fixed and stained with various antibodies as described previously (Hughes and Krause, 1999). The following primary antibodies were used at the indicated concentrations: mouse anti-FLAG (1:1000; Sigma); rat anti-HA (1:200; Roche); rat anti-Myosin (1:500; Abcam, Cambridge, MA); mouse anti-Myc (1:300; Cell Signaling); rabbit anti-Dmef2 (1:1000; from B. Paterson, National Cancer Institute, Bethesda, MD); rabbit anti-Tinman (1:1000); mouse anti-βPS-integrin (developed by Danny Brower and obtained from the Developmental Studies Hybridoma Bank, The University of Iowa, Department of Biological Sciences, Iowa City, IA, 1:500); mouse anti-β-Gal (Promega, 1:500). Alexa488-, Alexa568-, Alexa594-, and Alexa647-conjugatged secondary antibodies (Invitrogen) were used to recognize the primary antibodies. Muscle actin was stained by Alexa546-conjugated phalloidin (Invitrogen, 1:25). Images were obtained using a spinning disk confocal system (Ultraview ERS; Perkin Elmer, Norwalk, CT) mated to a CS9100-50; camera (Hamamatsu, Bridgewater, NJ), and an Axiovert 200M microscope (Carl Zeiss MicroImaging, Thornwood, NY) using Ultraview ERS software (Version 2, Perkin Elmer) and assembled with Adobe Photoshop (Ver. CS, San Jose, CA; using Windows XP, Microsoft, Redmond, WA).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Deshpande et al., 1997; Varadarajan and VijayRaghavan, 1999; Muller et al., 2005). Similar to what we observe with FISH detection of the sd signal, in sdETX4 embryos, significant levels of β-gal can be detected in some muscle cells (Figure 2A). This includes cardiac cells at stage 13 (data not shown), and by stage 16 β-gal is expressed in almost all cardiac cells in the heart region of the dorsal vessel (Figure 2A).
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). An advantage to this GAL4 based reporter is that we could use it to induce expression of a UAS-3xFLAG-Sd transgene. This allows the examination of a FLAG-tagged Sd under the control of an endogenous sd enhancer, closely mirroring the cellular produrance of the endogenous Sd protein. This consideration is important as sd expression driven by sd-GAL4 appears to be very dynamic. At stage 13, it drives expression in 31% cells of SMs and several cardiac cells (Figure 2, B and C). At early stage 16, it drives expression in all SMs but not in heart cells (Figure 2D). The expression appears to drop during late stage 16 and is restricted to only some ventral SMs (Figure 2E). It appears that sd-GAL4 induced expression of 3xFLAG-sd does not cause any dominant-negative changes to the somatic or cardiac muscle specification in embryos (Figure 2, B–F). We also analyzed the expression of UAS-β-gal driven by sd-GAL4. The β-Gal protein is extremely stable, which although makes interpretation of the dynamic nature of sd-GAL4 expression difficult in earlier stages, allowed us to confirm that expression is restricted to ventral SMs, the same as that of 3xFLAG-sd at late stage 16 (data not shown).
vg Is Expressed in Embryonic SMs But Not Heart Muscle
We used an anti-Vg antibody to correlate vg expression with that of the sd-reporters in embryonic muscles (Figure 2, C–E). 3xFLAG-Sd expression does not affect the expression of vg in muscle cells, because vg has the same expression pattern as in wild type. Vg is first detected at stage 11 in the progenitors of ventral SMs, VL1-4 (data not shown; see Figure 2 for the diagram of each muscle identity). Then it is present in the muscles, LL1 and DA1-3, at stage 13 (Figure 2C). Vg is also present in VL1-4, LL1, and DA1-3 when 3xFLAG-Sd appears in all SMs of early stage 16 embryos (Figure 2D) and when the expression of 3xFLAG-Sd fades and is restricted to some ventral SMs at late stage 16 (Figure 2E). After late stage 17, Vg cannot be reliably detected in muscle cells, confirming what has been reported previously (Baylies et al., 1998).
Both the sd3L and vgnull Mutations Cause Defects in Embryonic Muscle Development
The X-linked, recessive, sd3L allele is homozygous lethal. Sequencing of sd3L identified a T-A substitution producing a premature stop codon (Srivastava et al., 2004). The sd3L likely represents a strong loss-of-protein-function allele because some hemizygous male animals do hatch and survive as feeble larvae with behavioral abnormalities that maybe result from muscle defects (Campbell et al., 1991). For example, recently hatched wild-type larvae have characteristic contraction waves that pass from the posterior to anterior and are responsible for locomotion. We found that the waves of contraction are much slower in sd3L hemizygotes, taking approximately three times as long to pass from posterior tip to anterior tip compared with wild type. Notably, examination of the embryonic muscles of sd3L hemizygotes revealed defects in both heart and somatic muscle development (Figure 3, A–G). Many of these embryos (30%) have less than the wild-type number of cardiac cells (Figure 3C), and many of the remaining cardiac cells have nuclei larger than normal (Figure 3, A and B). In many of the mutant embryos we also see somatic muscle defects, most often the ventral SMs (VO4-6) get lost or have defects in development (Figure 3, D and E). Actin staining also revealed that the VO4-6 muscles disappeared in some segments (Figure 3, F and G).
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). vgnull larvae showed similar muscle contraction defects to those associated with sd3L, taking approximately two times as long for the contraction waves to reach the anterior tip compared with wild type. Actin staining showed that the VL2 muscle was often missing in vgnull embryos (Figure 3, F and H) with no detectable defects in other muscles. This phenotype is enhanced in Dmef2 RNA interference (RNAi) background (Supplemental Figure 2F), indicating a functional interaction between these two genes.
Sd, Vg, and Dmef2 Can Form a Multiprotein Complex
As some interaction of the mammalian homologues of Sd, Vg, and Dmef2 has been reported previously, coimmunoprecipitation (coIP) was performed from S2 cell lysates expressing epitope-tagged Vg, Sd, or Dmef2. These three proteins appear to form a tripartite complex as any two could be coIPed with the third (Figure 4A). For example, coIPs of 3xFLAG-Sd also could detect 3xHA-Vg and 6xMyc-Dmef2 (Figure 4A). Similar results were observed when we used 3xFLAG-Vg or 3xFLAG-Dmef2 to coIP the other two proteins (data not shown). The interactions between any two of these three proteins appear to be independent of the third, as coIP of any two does not require the coexpression of the third (Figure 4, B and D). To further test for the possibility that Vg is required for the interaction between Sd and Dmef2, sequential IPs were performed by first isolating 3xFLAG-Sd and 6xMyc-Dmef2 or 3xFLAG-Dmef2 and 6xMyc-Sd and then testing for the presence of Vg. In either case, Vg was not detected (Figure 4C). Time-course IPs were also performed to test the specificity of the interaction between Vg and Dmef2 (Figure 4D). This interaction appears to be highly specific, because the amount of 6xMyc-Dmef2 IPed by 3xFLAG-Vg increases with time.
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) were performed. We first confirmed that Vg can bind directly with Dmef2 (Figure 5A). We then tested deleted or truncated forms of Vg and found that only one Vg deletion, Vg3-9, did not interact with Dmef2 (Figure 5, B–D). This indicates that at least two independent domains within Vg participating in binding to Dmef2: one within amino acids 1-187 and the other within amino acids 279-336 (Figure 5C). The Sd-interaction domain of Vg has been mapped to amino acids 279–336 (Simmonds et al., 1998). We note that there is some residual binding of one of the deletions (Vg 1-4) with the GST control. However, the much stronger signal obtained when GST-Dmef2 is present makes us confident that this residual binding is due to weak, nonspecific "stickyness" of the N-terminal region of Vg. Because Vg8-9, a deletion that removes the Sd interacting region, still interacts strongly with Dmef2 (Figure 5B), Vg could potentially interact with Sd and Demf2 at the same time via different domains.
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). In the somatic muscles, expression of a known Dmef2 target, myosin is affected when Sd is present, and this effect is enhanced when both Vg and Sd are present (Figure 6, D–F). This phenotype is similar to that of embryos overexpressing a Dmef2 inhibitor, Him in SMs (Liotta et al., 2007). We observed a similar reduction in actin staining when Vg and Sd were overexpressed in cells also expressing Dmef2 (Figure 6, G–H). We then tested the mRNA levels of two known Dmef2 target genes, actin57B (act57B) and myosin heavy chain (mhc). Drosophila Act57B is the major myofibrillar actin expressed in skeletal, visceral, and cardiac muscle during embryogenesis, and there is a conserved Dmef2-binding site within the promoter that is necessary for the full expression of act57B in embryos (Kelly et al., 2002). Similarly, mhc expression is significantly reduced in Dmef2 mutants (Lilly et al., 1995). When elevated levels of Sd and Vg are present in Dmef2-expressing cells, a significant reduction in both act57B and mhc mRNA are observed (Figure 6I). To confirm that the effect we see on suppression of actin or myosin is specific to the presence of Vg and Sd, we also expressed these transgenes in a single muscle using a Gal4 driver specific for VL1 (Figure 6K). We see a corresponding reduction in actin or myosin expression in the VL1 muscle with increased expression of sd and vg (Figure 6, J and K). These reductions lead to the missing of myofiber in SMs (Figure 6, H and K). Altogether, these data indicate that Sd and Vg repress Dmef2 function during muscle differentiation.
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; Lilly et al., 1995; Maeda et al., 2002a), we used Dmef2-GAL4 to express transgene(s) in all mesodermal cells of late stage 7 embryos and later in cardiac cells of the heart, visceral, and somatic muscle cells until the end of embryogenesis (Ranganayakulu et al., 1998). The pattern of the muscle system in wild-type embryos is shown in Figure 7, A and B. An act57B-LacZ transgene was used to visualize muscle fibers (Figure 7A) and to monitor for gross alterations in the expression level of act57B in muscle cells.
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We further examined the organization of the developing muscles by staining for βPS-integrin, one of the major integrins, acting as a transmembrane protein that stabilizes attachments between two neighbor muscles and that between muscles and epidermis along the segment border (Volk and VijayRaghavan, 1994; Brown et al., 2000). In wild-type embryos, muscle cells attach at characteristic positions relative to segment borders (Figure 7H). However, in embryos expressing UAS-3xFLAG-sd and UAS-3xHA-vg, the organization of muscle cell attachment is severely disrupted (Figure 7I). Embryos ectopically expressing UAS-3xHA-vg in SMs exhibit a different phenotype. SMs still have a highly organized pattern (Figure 7C), and muscle cells do not lose their positions (data not shown), but the migration of VO4-6 muscles seem to be inhibited or redirected (Figure 7, F and G). However, expression of UAS-vg 3-9, the Vg deletion that loses interaction with Dmef2 (Figure 5C), produced wild-type phenotype (data not shown). As a control, we examined embryos overexpressing UAS-6xMyc-Dmef2, and this does not cause any obvious defect in SMs (data not shown).
Altered Expression of Sd, Vg, and Dmef2 Causes Defects in Cardiac Cell Development
In wild-type, there is a single row of Dmef2-positive cardiac cells on each side of the embryo (Figure 8A) and four Tin-positive cardiac cells per hemisegment with some Tin-positive pericardial cells (Figure 8B, see Figure 2F for a diagram of each cardiac cell fate). It was reported previously that the Dmef2-GAL4 driver was not active in pericardial cells (Ranganayakulu et al., 1998), but, based on our analysis of the pattern of β-gal expression driven by Dmef2-GAL4, it also drives expression, at least transiently, in pericardial cells (Figure 8A'). We used this driver for examining the effect of elevated levels of Vg or Sd on cardiac cell differentiation.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Shyamala and Chopra, 1999) match the Sd expression pattern in embryonic muscles revealed by sd in situ (Figure 1). We have also determined that Vg, the first known TEF-1 family cofactor, has a role in SMs but not cardiac muscles (Figure 3H), and in fact, elevating Vg expression in the developing dorsal vessel has a negative effect on differentiation of this organ. Our results show several lines of evidence supporting a model whereby Sd and Vg, in a complex with Dmef2, help to regulate late-stage Drosophila embryonic muscle development. One prediction of this model is that altering the relative levels of these proteins will have significant effects on specific subsets of muscle cells. The effect of the relative levels of each of these proteins can be observed in various cell types: 1) Sd and Dmef2 are coexpressed in the cardiac cells (Figure 2, A and B), where vg is not expressed; 2) Starting at stage 11, vg and Dmef2 are coexpressed in the progenitors of some SMs (Figure 2, C–E), where sd is expressed at a later stage (Figure 2, C–E); and 3) These three genes are coexpressed in SMs, DA1-3, LL-1, and VL1-4 at early stage 16 (Figure 2D), but by late stage 16 the coexpression is restricted to some ventral SMs (Figure 2E). We have also shown that 1) Sd is able to interact with Dmef2 without the presence of Vg (Figure 4, B and C); 2) Vg is able to interact with Dmef2 without the presence of Sd (Figure 4D); and 3) It is also possible for Sd, Vg and Dmef2 to form a tripartite complex (Figure 4A). Given that Vg appears to bind Dmef2 at two different sites, it may be that Vg could be the bridge protein connecting Sd and Dmef2, because Vg can bind each of them via a different domain (Figure 5, B and C). Finally, a requirement for the presence of Sd and Vg appears to be specific to differentiation of specific muscle types as mutations in sd and vg cause defects in different muscles (Figure 3), whereas alterations in the relative expression levels of any of these three genes in developing muscles of Drosophila caused specific alterations in both SM and cardiomyoctes (Figures 7 and 8).
Because our data show that Vg can bind Dmef2 independently of Sd, it is possible that Vg may modify Dmef2 activity in the absence of Sd. We noted that the vg and Dmef2 genes are coexpressed in some SMs (DA1-3, LL1, and VL1-4) before Sd is present in those muscles (Figure 2, C–E). They are also coexpressed in the progenitors of muscle VL1-4. We also show that functional interactions exist between Vg and Dmef2, as coexpression of them in heart cells has a synergistic effect on increasing the numbers of Dmef2-positive and Tin-positive cardiac cells (Figure 8, I and J), and the phenotype of vgnull mutant is enhanced in a Dmef2 deficiency background (Supplemental Figure S2F). Previous studies of vg have almost exclusively focused on its function as a wing identity gene. However, there is now mounting evidence that Vg also defines the cellular identity of a subgroup of embryonic SMs (Figures 3 and 7, C and G; Baylies et al., 1998; Sudarsan et al., 2001), although the functional role of Vg in the development of these muscles is not clear. Dmef2 is considered to be a "differentiation gene" playing a role in the final stages of muscle differentiation. Thus, this begs the question: what is the significance of the interaction between these two proteins that apparently have roles at different developmental stages? A recent study showed that Dmef2 not only binds to regulatory regions of muscle structural genes but also binds many muscle "identity genes" and genes involved in early signal pathways of muscle development (Sandmann et al., 2006), indicating a role of Dmef2 in early muscle development. Therefore, Vg may act together with Dmef2 to specify those SMs in which Vg is expressed. Our data support this idea, because vgnull mutants often lose muscle VL-2 (Figure 3H), and overexpression of Sd leads to either poor development or loss of muscle LL1 and VL1-4 (Figure 7, D1–D3) where Vg is present. Considering the strong functional interaction that is known to occur between Vg and Sd (Simmonds et al., 1998), overexpression of Sd may interfere with the function of Vg in those muscles.
Just as Vg and Dmef2 may interact in the absence of Sd, a Sd/Dmef2 complex may exist in muscle cells where vg is not expressed significantly; i.e., cardiac cells in the heart region and some somatic muscle cells (Figure 2). Expression of UAS-6Myc-Dmef2 via Dmef2-GAL4 results in one extra row of Dmef2-positive cardiac cells (Figure 8G). This phenotype is not unexpected as Dmef2-GAL4 is also active in pericardial cells that surround cardiac cells (Figure 8A'). However, it is unexpected that expression of UAS-3xFlAG-sd also produces extra rows of Dmef2-positive cardiac cells (Figure 8E). These results indicate that Sd could activate the expression of Dmef2 in the pericardial cells. Because the pattern of expression directed by the enhancer of Dmef2 in muscle cells is very complicated (Nguyen and Xu, 1998) it has been proposed that there is an autoregulation mechanism to maintain its expression in differentiated muscles (Cripps et al., 2004). Therefore, Sd might be required to act with Dmef2 to maintain expression of Dmef2 in cardiac cells at late stages. The ability of Dmef2 to partially rescue the heart phenotype caused by expression of UAS-3xFLAG-sd (Figure 8, K–M) also suggests a functional interaction between Sd and Dmef2, because Sd itself does not have transcriptional activation ability and overexpression of Sd can lead to repression of transcription (Simmonds et al., 1998).
Mutation and ectopic-expression analysis also revealed that Sd has a role in both heart muscle and SM development (Figure 3). Recently, Sd was shown to be the target of the Hippo (Hpo) signaling pathway that governs cell growth, proliferation, and apoptosis (Zhang et al., 2008). Inactivation of Sd diminishes Hpo target gene expression and reduces organ size, whereas a constitutively active Sd promotes tissue overgrowth (Zhang et al., 2008). We see that in sd3L mutants there are fewer heart cells and that the VO4-6 muscles appear to have defects in their differentiation (Figure 3, A–F), whereas overexpression of Sd in VO4-6 produces more projections (Figure 7D). These phenotypes would suggest a role of Sd in both growth and proliferation of muscle cells. Conversely, ectopic expression of Vg in VO4-6 muscles leads to a phenotype similar to that of sd3L (Figure 7, F and G). Thus, it appears that ectopic expression of Vg in those muscles interferes with the function of Sd.
We observed that coexpression of UAS-3xFLAG-sd and UAS-3xHA-vg for extended times via Dmef2-GAL4 causes significant defects in muscle differentiation, including significant alterations in their sites of attachment. In cardiac muscles, Tin-positive heart cells end up in the SM region (Figure 8O); by the end of muscle development, the stereotyped patterning of SMs is totally disrupted (Figure 7E). This phenotype may be a result of the apparent dynamic expression we observe of the sd reporters in SMs (Figure 2). Thus, any Sd–Vg complex that would be formed in developing muscles would be transient, freeing each potential cofactor to interact with Dmef2 independently.
It is interesting that Sd–Vg complex represses Dmef2 function without affecting Dmef2 expression during muscle development (Figure 6 and Supplemental Figure S2E). The protein Him (Holes in muscle) was also shown before to repress Dmef2 function during muscle differentiation, and the authors argue that a balance of positive and negative inputs controls muscle differentiation (Liotta et al., 2007). Our data support this idea and may reveal another layer of negative input, the Sd–Vg complex, in muscle differentiation, because overexpression of Sd or Sd and Vg produces a phenotype similar to that of overexpression of Him in developing SMs (Figure 6, E and F) and also to that of Dmef2 RNAi embryos. The repression we see of act57B, the product of which is primarily required during muscle differentiation, may be a normal occurrence during late stage 16 when most SMs are presumably fully differentiated, having finished migration and reached their attachment sites (Schnorrer and Dickson, 2004). At this time, some SMs in different segments contact each other and specific extracellular matrix (ECM) contacts between muscles form (Martin-Bermudo, 2000; Martin-Bermudo and Brown, 2000). act57B is initially expressed in SMs at stage11, and by stage16 there is already high levels of act57B transcript in SMs (Kelly et al., 2002). Specifically, these SMs would slow myofibril growth by repressing the expression of act57B, especially in those SMs that contact with neighboring muscles, like LL1 and VL1-4, and the presence of Vg in these muscles (Figure 3H) may be mediating this repression.
The most significant repression of Dmef2 function appears to require the presence of both Sd and Vg (Figure 6F and Supplemental Figure S2D). However, this is at odds with the presumptive activating function mediated by an Sd–Vg complex that occurs in other tissues like the wing imaginal disk, where an Sd–Vg complex binds and activates the vg boundary enhancer (Halder et al., 1998). The differential activities of these proteins in muscle versus wing development may reflect a requirement for yet additional proteins within a presumptive Vg/Sd/Dmef2 complex to modify its activity in a tissue-specific manner. Alternatively, posttranslational modifications to Vg or Sd (or both) may modify their activity. Interestingly, the yeast Sd homologue, Tec1, is phosphorylated and then degraded during the mating pheromone response (Bao et al., 2004). In mammals, TEF-1 is phosphorylated responding to cAMP/PK-A signaling (Gupta et al., 2000). However, there is, as yet, no clear indication that Sd is phosphorylated in Drosophila cells.
Finally, although we have shown that Sd, Vg, and Dmef2 interact directly, similar to their mammalian homologues, our data suggest potential new functions of Sd and Vg during muscle specification. For example, Vg seems to have role in the specification of ventral muscles VL1-4, and Sd has a role in the development of muscle VO4-6, especially in the development of their projections. In addition, the Sd–Vg complex represses Dmef2 function, which is at odds with the known activities of their mammalian homologues. However, this repression only happens in certain muscles (e.g., VL1-4) that need to contact neighboring muscles.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Andrew J. Simmonds (andrew.simmonds{at}ualberta.ca).
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REFERENCES |
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31% cells of somatic muscles (SMs, arrowheads) at stage 13. Note that sd-GAL4 is also activated in cells of central neuron system (CNS, empty arrow). Dmef2 is present in all muscle cells. C1–C3 shows the dorsal SMs where vg is expressed at stage 13. 3xFLAG-Sd can be detected in some SMs. D1–D3 shows that vg is expressed in the DA3, LL1, and VL1-4 muscles when 3xFLAG-Sd appears in all SMs at stage 16. Vg also appears in some neuronal cells (arrowheads). DA1-2 are not shown because they are out of the field of view. E1–E3 shows that vg is still expressed in the DA3, LL1, and VL1-4 muscles when the expression of 3xFLAG-Sd fades in SMs and appears only in some ventral SMs at late stage 16. At this stage, 3xFLAG-Sd appears with Vg in the neuron cells shown above (arrowheads). (F) A schematic drawing of a stage 16 embryonic dorsal vessel (dorsal view, anterior to the left). Heart cells include two parallel rows of Dmef2-positive cardiac cells in the middle with four Tinman-positive cardiac cells per hemisegment starting from T1. Dmef2-negative pericardial cells surround the cardiac cells. On the right is a schematic representation of the embryonic SMs in each abdominal hemisegment A2–A7 (lateral view with anterior up) using the nomenclature of Crossley (1978)
