Myocyte enhancer factor 2



Mef2 gene expression is first detected during Drosophila embryogenesis within mesodermal precursor cells during gastrulation [Images], prior to specification of the somatic and visceral muscle lineages (Lilly, 1994). MEF2 protein is expressed throughout the mesoderm following gastrulation. Later in embryogenesis, its expression is maintained in precursors and differentiated cells of the somatic and visceral musculature, as well as the heart. Like Nautilus, Mef2 seems to be confined to single precursor cells.

The three major types of musculature in the Drosophila embryo requireMEF2 for later aspects of their differentiation, including body wall muscles, gut musculature, and the heart . tinman is restricted to the visceral mesoderm, the tissue that gives rise to cardiac muscle cells, and nautilus is restricted to somatic mesoderm, the tissue that gives rise to skeletal cells, however MEF2 is found in both tissues. (Bour, 1995).

In Drosophila, much has been learned about the specification of neuronal cell fates but little is known about the lineage of mesodermal cells with different developmental fates. During development, individual mesodermal precursor cells are initially singled out to become the founder cells for specific muscles. The selection of muscle founder cells is thought to employ a Notch-mediated process of lateral inhibition, similar to what is observed for the specification of neural precursors. These muscle founder cells then seem to fuse with the surrounding, uncommitted myocytes, inducing the formation of muscle fiber syncytia. In contrast, the differentiated progeny of neural precursor cells are usually the result of a fixed pattern of asymmetric cell divisions that are directed, in part, by interactions among Numb (a localized intracellular-receptor protein); Sanpodo (Spdo, a potential tropomodulin homolog), and Notch (a transmembrane receptor protein). The roles of these neural lineage genes have been examined in the cell fate specification of muscle and heart precursors. numb and spdo mutations have opposite effects in the specification of muscle founder cells. In all numb mutant embryos examined, the number of Kruppel and S59/NK1 expressing muscle cells is dramatically reduced or absent in stage 12/13 embryos. In spdo mutant embryos the number of S59 and Kr expressing muscle founders is increased (Park, 1998).

A progenitor cell that generates both a pericardial heart cell and a somatic muscle founder cell was the focus of investigation. The two sibling cells studied were a single dorsal muscle, DA1, and a non-muscle pericardial cell (termed EPC), which is associated with the heart. Both cell types express Eve, however, they can be distinguished from one another morphologically. The precursor for both the EPCs and the putative founders of DA1 muscle emerges from a small cluster of mesodermal Eve-expressing cells in each hemisegment at mid-stage 11. At first, these mesodermal Eve cells are indistinguishable from one another, and they co-express Mef2, which is expressed in the entire early mesoderm and later in all (contractile) muscle types. Subsequently Mef2 expression ceases in the future EPCs as they begin to differentiate as non-muscle, pericardial cells. The putative DA1 founder seems to maintain Mef2 expression and begins fusing with surrounding myocytes. In Mef2 mutants, no fusion occurs but the putative muscle founders maintain expression of their muscle precursor markers, such as Eve. The asymmetric segregation of Numb into one of these daughter cells antagonizes the function of Notch and Spdo by preventing the presumptive muscle founder from assuming the same fate as its cardiac sibling. In numb mutants, most DA1 muscles are physically absent and the remaining ones lack Eve (and Kr) expression; in addition, the number of EPCs is doubled. These data suggest that in numb mutants, the putative DA1 founders are transformed into EPCs, because the Eve progenitor cells that normally give rise to DA1 founders and EPCs now only produce EPCs. Overexpression of numb or loss-of-spdo-function result in a failure to generate EPCs but allow for the formation of DA1 muscles. Similarly, expression of constitutively active Notch leads to a failure of DA1 muscle formation and an increase in the number of EPCs. Studies of double mutants indicate that spdo is epistatic to numb (in numb;spdo double mutant embryos, the spdo phenotype is apparent), suggesting that it acts downstream of numb. These results suggest that asymmetric cell divisions, in addition to the previously-documented inductive mechanisms, play a major role in cardiac and somatic muscle patterning. In addition, the cytoskeleton may have a role in the asymmetrical localization of cell fate determinants (Park, 1998).

Subdivision and developmental fate of the head mesoderm in Drosophila

This paper defines temporal and spatial subdivisions of the embryonic head mesoderm and describes the fate of the main lineages derived from this tissue. During gastrulation, only a fraction of the head mesoderm (primary head mesoderm; PHM) invaginates as the anterior part of the ventral furrow. The PHM can be subdivided into four linearly arranged domains, based on the expression of different combinations of genetic markers (tinman, heartless, snail, serpent, mef-2, zfh-1). The anterior domain (PHMA) produces a variety of cell types, among them the neuroendocrine gland (corpus cardiacum). PHMB, forming much of the'T-bar' of the ventral furrow, migrates anteriorly and dorsally and gives rise to the dorsal pharyngeal musculature. PHMC is located behind the T-bar and forms part of the anterior endoderm, besides contributing to hemocytes. The most posterior domain, PHMD, belongs to the anterior gnathal segments and gives rise to a few somatic muscles, but also to hemocytes. The procephalic region flanking the ventral furrow also contributes to head mesoderm (secondary head mesoderm, SHM) that segregates from the surface after the ventral furrow has invaginated, indicating that gastrulation in the procephalon is much more protracted than in the trunk. This study distinguishes between an early SHM (eSHM) that is located on either side of the anterior endoderm and is the major source of hemocytes, including crystal cells. The eSHM is followed by the late SHM (lSHM), which consists of an anterior and posterior component (lSHMa, lSHMp). The lSHMa, flanking the stomodeum anteriorly and laterally, produces the visceral musculature of the esophagus, as well as a population of tinman-positive cells that is interpreted as a rudimentary cephalic aorta ('cephalic vascular rudiment'). The lSHM contributes hemocytes, as well as the nephrocytes forming the subesophageal body, also called garland cells (de Velasco, 2005).

The mesoderm is a morphologically distinct cell layer that can be recognized in early embryos of most bilaterian phyla and that gives rise to tissues interposed between ectodermal and endodermal epithelia, including muscle, connective, blood, vascular, and excretory tissue. Besides the differentiative fate of tissues derived from it, the mesoderm shares several common properties in regard to its formation during gastrulation. The anlage of the mesoderm is sandwiched in between the anlage of the endoderm and the neurectoderm. This has been documented in most detail in anamniote vertebrates, where signals from the vegetal blastomeres (the anlage of the endoderm) act on the adjacent marginal zone of the future ectoderm to induce mesoderm. Although gastrulation proceeds quite differently in arthropods from the way it does in chordates, the proximity of the mesodermal anlage to future endoderm and neurectoderm is conserved, and numerous signaling pathways and transcriptional regulators that share similar function and expression patterns in arthropods and chordates have been identified (de Velasco, 2005 and references therein).

Following gastrulation, the mesoderm is subdivided along the dorso-ventral axis into several subdivisions laid out in a distinct dorso-ventral order. In vertebrates, cells located in the dorsal part of the mesoderm anlage give rise to notochord and somites, which in turn produce muscular, skeletal, and connective tissue. Next to the somitic mesoderm is the intermediate mesoderm that will form the excretory and reproductive system. The ventral mesoderm (lateral plate) gives rise to blood, vascular system, visceral musculature, and coelomic cavity. In arthropods, fundamentally similar mesodermal subdivisions can be recognized, and similarities extend to the relative positions these domains obtain relative to each other and relative to the adjacent neurectoderm. For example, precursors of visceral muscles, vascular system, and blood are at the edge of the mesoderm facing away from the neural primordium (ventral in vertebrates, dorsal in arthropods (de Velasco, 2005 and references therein).

The subdivision of the vertebrate mesoderm into distinct longitudinal tissue columns with different fates is seen throughout the trunk and head of the embryo. However, several significant differences between the head and the trunk are immediately apparent. For example, cells derived from the anterior neurectoderm form the neural crest that migrates laterally and gives rise to many of the tissues that are produced by mesoderm in the trunk. As a result, the fates taken over by the head mesoderm are more limited than those of the trunk mesoderm. In contrast, the head mesoderm produces several unique lineages, such as the heart (cardiac mesoderm) and a population of early differentiating macrophages. Moreover, some of the signaling pathways responsible for inducing different mesodermal fates in the trunk appear to operate in a different manner in the head. A recently described example is the Wnt signal that induces somatic musculature in the trunk, but inhibits the same fate in the head (de Velasco, 2005 and references therein).

The head mesoderm of arthropods, like that of vertebrates, also appears to deviate in many ways from the trunk mesoderm. For example, specialized lineages like embryonic blood cells and nephrocytes forming the subesophageal body (also called garland cells) arise exclusively in the head. That being said, very little is known about how the arthropod head mesoderm arises and what types of tissues derive from it. The existing literature mainly uses histology, which severely limits the possibilities of following different cell types forward or backward in time. In this paper, several molecular markers have been used to initiate more detailed studies of the head mesoderm in Drosophila. The goal was to establish temporal and spatial subdivisions of the head mesoderm and, using molecular markers expressed from early stages onward, to follow the fate of the lineages derived from this embryonic tissue. Besides hemocytes and pharyngeal muscles described earlier, the head mesoderm also gives rise to several other lineages, including visceral muscle, putative vascular cells, nephrocytes, and neuroendocrine cells. The development of the head mesoderm is discussed in comparison with the trunk mesoderm and in the broader context of insect embryology (de Velasco, 2005).

The Drosophila head mesoderm, as traditionally defined, includes all mesoderm cells originating anterior to the cephalic furrow. The formation of the head mesoderm is complicated by the fact that (unlike the mesoderm of the trunk) only part of it invaginates with the ventral furrow; by far, the majority of head mesoderm cells, recognizable in a stage 10 or 11 embryo, segregate from the surface epithelium of the head after the ventral furrow has formed. Another complicating factor is that head mesoderm cells derived from different antero-posterior levels adopt very different fates, unlike the situation in the trunk where mesodermal fates within different segments along the AP axis are fairly homogenous, with obvious exceptions such as the gonadal mesoderm that is derived exclusively from a subset of abdominal segments. Using several different markers, this study has followed the origin, migration pathways, and later, fates of head mesoderm cells (de Velasco, 2005).

The anterior part of the ventral furrow, called primary head mesoderm (PHM) in the following, includes cells that will contribute to diverse tissues, including muscle, hemocytes, endoderm, and several ill-defined cell populations closely associated with the brain and neuroendocrine system. For clarification, the anterior ventral furrow will be divided into the following domains:

The anterior lip of the T-bar (PHMA) is the source of the corpus cardiacum, as well as other gt-positive cells that at least in part end up as nerve cells flanking the frontal connective and frontal ganglion. These cells continue the expression of giant throughout late embryonic development; they represent a hitherto unknown class of nonneuroblast-derived neurons (de Velasco, 2005).

The posterior lip of the T-bar (PHMB) can be followed towards later stages by its continued expression of htl. These cells, called the procephalic somatic mesoderm, form a bilateral cluster that moves dorso-anteriorly into the labrum and becomes the dorsal pharyngeal musculature. Htl expression almost disappears in these cells around late stage 11, but is reinitiated at stage 12 and stays strong until stage 14, when the dorsal pharyngeal muscles differentiate. Many of the genes expressed in the somatic musculature of the trunk and its precursors (Dmef2, beta-3-tubulin) are also expressed in the procephalic somatic mesoderm (de Velasco, 2005).

The part of the ventral furrow posteriorly adjacent to the T-bar (PHMC) expresses srp, forkhead (fkh), and other endoderm/hemocyte markers. After the ventral furrow closes in the ventral midline (stage 7/8), these cells form a compact median mass, most of which represents part of the anterior endoderm that gives rise to the midgut epithelium. Starting at around this stage, the lateral part of the hemocyte-forming 'secondary head mesoderm' ingresses in between the endoderm and the surface ectoderm. It is likely that some of the PHMC cells invaginating already with the ventral furrow, along with the cells that form the anterior endoderm, also give rise to hemocytes. Precursors of hemocytes and midgut are difficult to distinguish during and shortly after ventral furrow invagination since both express srp and other markers shared between hemocytes and midgut precursors. At around stage 9, the two populations of precursors disengage. The endoderm remains a compact mesenchyme attached to the invaginating stomodeum; hemocyte precursors move dorsally and take on the shape of expanding vertical plates interposed in between endoderm and ectoderm (de Velasco, 2005).

Domain PHMD, the short portion of the ventral furrow situated posterior to the endoderm, along with a considerable portion of the mesoderm behind the cephalic furrow, forms the mesoderm of the three gnathal segments (mandible, maxilla, labium). The gnathal mesoderm in many ways behaves like the mesoderm of thoracic and abdominal segments. It gives rise to somatic muscle (the lateral pharyngeal muscles), visceral muscle, and fat body. Unlike trunk mesoderm, gnathal mesoderm does not produce cardioblasts and pericardial cells. Instead, a large proportion of gnathal mesoderm cells, joining the anteriorly adjacent secondary procephalic mesoderm, adopt the fate of hemocytes (de Velasco, 2005).

Besides the ventral furrow, other parts of the ventral procephalon produce head mesoderm in a complex succession of delamination and ingression events. The head mesoderm that forms from outside the ventral furrow will be called 'secondary mesoderm' (SHM) in the following. Based on the time of formation and the position relative to the stomodeum, the following phases and domains of secondary head mesoderm development can be distinguished.

Following the obliteration of the ventral furrow at stage 8, the eSHM delaminates from the ventral surface 'meso-ectoderm' (considering that this epithelium still contains mesodermal progenitors!) flanking the endodermal mass. The eSHM forms two monolayered sheets that gradually move dorsally and posteriorly; by stage 9, the eSHM cells line the basal surface of the emerging head neuroblasts. An undefined number of primary head mesoderm cells derived from domain PHMC of the ventral furrow are mingled together with the eSHM cells. The ultimate fate of the eSHM is that of hemocytes: they express srp, followed slightly later by other blood cell markers (e.g., peroxidasin and asrij). A subset of hemocytes, called crystal cells, derive from precursors that form a morphologically conspicuous cluster at the dorsal edge of the eSHM, identifiable from early stage 10 onward by the expression of lz. The mechanism by which at least part of the eSHM delaminates is unique. Thus, it is formed by the vertically oriented division of the surface epithelium, whereby the inner daughters will become eSHMe and the outer ones ectoderm. The focus of vertical mitosis has named the procephalic domain in which it occurs 'mitotic domain #9' (de Velasco, 2005).

From late stage 9 onward, the early SHMs are followed inside the embryo by the closely adjacent posterior late SHMs. One cluster of posterior late secondary head mesoderm (lSHMp) cells delaminates from the surface epithelium flanking the posterior lip of the stomodeum; a second lSHMp cluster appears at the same stage at a slightly more posterior level. The first cluster seems to contribute to the hemocyte population; the posterior cluster gives rise to the nephrocytes forming the subesophageal body (also called garland cells; labeled by CG32094). Garland cell precursors are initially arranged as a paired cluster latero-ventrally of the esophagus primordium; subsequently, the clusters fuse in the midline and form a crescent underneath the esophagus. Garland cells are distinguished from crystal cells by their size, location, and arrangement: crystal cells are large, round cells grouped in an oblong cloud dorso-anterior to the proventriculus. Garland cells are smaller, closely attached to each other, and lie ventral of the esophagus (de Velasco, 2005).

During stages 10 and 11, cells delaminate beside and anterior to the stomodeum, originating from the anlage of the esophagus and the epipharynx (labrum). These cells, called anterior late secondary head mesoderm cells (lSHMa), can be followed by their expression of tin. Two groups can be distinguished. The tin-positive cells delaminating from the esophageal anlage (es) give rise to the visceral musculature (vm) surrounding the esophagus. These cells lose tin expression soon after their segregation, but can be recognized by other visceral mesoderm markers such as anti-Connectin. More dorsally, in the anlage of the clypeolabrum (cl) delaminate, the dorsal subpopulation of the lSHMas, which rapidly migrates posteriorly on either side and slightly dorsal of the esophagus, can be found. These cells retain expression of tin into the late embryo. They assemble into two longitudinal rows stretching alongside the roof of the esophagus primordium. During late embryogenesis, they move posteriorly along with the esophagus towards a position behind the brain commissure. Many of the tin-positive SHMs apparently undergo apoptosis: initially counting approximately 25 on either side, they decrease to 12-15 at stage 14 to finally form a single, irregular row of about 15 cells total in the late embryo. These cells come into contact with the anterior tip of the dorsal vessel. This formation of previously undescribed cells, for which the term 'procephalic vascular cells', is proposed, is interpreted as a rudiment of the head aorta, which forms a prominent part of the dorsal vessel in many insect groups (de Velasco, 2005).

On the basis of additional molecular markers, the tin-positive procephalic vascular cells are further subdivided into two populations. The first subpopulation expresses the muscle and cardioblast-specific marker Dmef2; the second type is Dmef2-negative. In the dorsal vessel of the trunk, tin-positive cells also fall into a Dmef2-positive and a Dmef2-negative population. Dmef2-positive cells of the trunk represent the cardioblasts, myoendothelial cells lining the lumen of the dorsal vessel. Dmef2-negative/tin-positive cells form a somewhat irregular double row of cells attached to the ventral wall of the dorsal vessel. The ultimate fate of these cells has not been explored yet. However, preliminary data suggest that they develop into a muscle band that runs alongside the larval dorsal vessel. This would correspond to the situation in other insects in which such a ventral cardiac muscle band has been described (de Velasco, 2005).

The role of tinman in the formation of the procephalic vascular rudiment was investigated by assaying tin-mutant embryos for the expression of Dmef2. Similar to the cardioblasts of the trunk, the Dmef2-positive cells of the procephalic vascular rudiment are absent in tin mutants. It is quite likely that the (Dmef2-negative) remainder of the procephalic vascular rudiment is affected as well by loss of tin, but in the absence of appropriate markers (besides tin itself, which is not expressed in the mutant), it was not possible to substantiate this proposal (de Velasco, 2005).

At the time of appearance of the ventral furrow, segmental markers such as hh do not allow the distinction between distinct 'preoral' segments. Thus, hh is expressed in a wide procephalic stripe in front of the regularly sized mandibular stripe. During stage 7, the procephalic hh stripe splits into an anterior, antennal stripe and a posterior, short, intercalary stripe. The anterior lip of the ventral furrow (domain PHMA) coincides with the anterior boundary of the antenno-intercalary stripe. Thus, the primary head mesoderm and endoderm originating from within the anterior ventral furrow can be considered a derivative of the antennal and intercalary segments. This interpretation is supported by the expression of the homeobox gene labial (lab) found in the intercalary segment. The labial domain covers much of the anterior ventral furrow, including domains PHMB-C (de Velasco, 2005).

Morphogenetic movements in the ventral head, associated with the closure of the ventral furrow, the formation of the stomodeal placode, and the subsequent invagination of the stomodeum result in a shift of head segmental boundaries. The antennal segment tilts backward, as can be seen from the orientation of the antennal hh stripe that from stage 8 onward forms an almost horizontal line, connecting the cephalic furrow with the sides of the stomodeal invagination (which falls within the ventral realm of the antennal segment, in Drosophila as well as other insects). Since the expression of hh, like that of engrailed (en), coincides with the posterior boundary of a segment, the territory located ventral to the antennal hh stripe falls within the intercalary segment. This implies that most, if not all, of the posterior late SHM, is intercalary in origin. It is further plausible to consider that the anterior lSHM belongs to the intercalary and antennal segment. The vascular cells of the head, a conspicuous derivative of the anterior lSHM in Drosophila, are derived from the antennal mesoderm in other insects. The labrum, with which much of the anterior lSHM is associated, represents a structure that has always been difficult to integrate in the segmental organization of the head. Most likely the labrum represents part of the intercalary segment; this would help explain some of the unusual characteristics of the head mesoderm (de Velasco, 2005).

In conclusion, several fundamental similarities are found between the mesoderm of the head and that of the trunk regarding the tissues they give rise to, and possibly the signaling pathways deciding over these fates. After an initial phase of structural and molecular homogeneity, the trunk mesoderm becomes subdivided into a dorsal and a ventral domain by a Dpp-signaling event that emanates from the dorsal ectoderm. The dorsal domain, characterized by the Dpp-dependent continued expression of tinman, becomes the source of visceral and cardiogenic mesoderm, among other cell types. A role of Dpp/BMP signaling in cardiogenesis seems to be conserved among insects and vertebrates. Subsequent signaling steps, involving both Wingless and Notch/Delta, separate between these two fates and further subdivide the cardiogenic mesoderm into several distinct lineages, such as cardioblast, pericardial cells, and secondary hemocyte precursors (lymph gland). As a result of these signaling events, Tinman and several other fate-determining transcription factors become restricted to their respective lineages: tin to the cardioblasts, odd to pericardial cells and hemocyte precursors, zfh1 and srp to hemocyte precursors and fat body. Dmef2 and several other transcription factors become restricted to various combinations of muscle types (somatic, visceral, cardiac) (de Velasco, 2005).

In the head mesoderm, the above genes are associated with similar fates. Tin and Dmef2 appear widely in the procephalic ventral furrow and the anterior lSHM before getting restricted to the procephalic vascular rudiment and/or the pharyngeal musculature, respectively. In contrast with the initially ubiquitous expression of Tin and Dmef2 in the trunk mesoderm, those parts of the head mesoderm giving rise to hemocytes (PHMC, posterior lSHM) never express these mesodermal genes. Previous work has shown that the head gap gene buttonhead (btd) is responsible for the early repression of tin in the above mentioned domains of the head mesoderm. The early absence of Tin and Dmef2 in the head mesodermal hemocyte precursors is paralleled by the presence of Srp and Zfh1 in these cells. Interestingly, Srp/Zfh-positive cells of the head produce only hemocytes and no fat body, suggesting that an as-yet-uncharacterized signaling step prevents the formation of fat body in the head. It is tempting to speculate that there exists within the mesoderm a 'blood/fat body equivalence group'. Blood cells and fat body share not only the expression of fate-determining genes such as srp and zfh1, but also, later, functional properties that have to do with immunity. In the trunk, the blood/fat body equivalence group gives rise mostly to fat body, producing only a limited number of hemocyte precursors in the dorsal mesoderm of the thoracic segments. In the head, on the other hand, all cells of the equivalence group become hemocytes (de Velasco, 2005).

Attention is drawn to another mesodermal lineage that produces related, yet not identical, cell types in the trunk and the head: the nephrocytes. Nephrocytes are defined by their characteristic ultrastructure (membrane invaginations sealed off by junctions) that attests to their excretory function. In the trunk, nephrocytes are represented by the pericardial cells that settle beside the cardioblasts; a newly discovered nephrocyte population ('star cells') invading the Malpighian tubules is derived from the mesoderm of the tail segments. In the head, nephrocytes aggregate near the junction between esophagus and proventriculus as the subesophageal body, also called garland cells. The fact that from the early stages of development onward different transcription factors are expressed in garland cells and pericardial cells suggests that these cells perform similar, yet not fully overlapping, functions (de Velasco, 2005).


Drosophila metamorphosis is characterized by diverse developmental phenomena, including cellular proliferation, tissue remodeling, cell migration, and programmed cell death. Cells undergo one or more of these processes in response to the hormone 20-hydroxyecdysone (ecdysone), which initiates metamorphosis at the end of the third larval instar and before puparium formation (PF) via a transcriptional hierarchy. Additional pulses of ecdysone further coordinate these processes during the prepupal and pupal phases of metamorphosis. Larval tissues such as the gut, salivary glands, and larval-specific muscles undergo programmed cell death and subsequent histolysis. The imaginal discs undergo physical restructuring and differentiation to form rudimentary adult appendages such as wings, legs, eyes, and antennae. Ecdysone also triggers neuronal remodeling in the central nervous system (White, 1999).

Wild-type patterns of gene expression in D. melanogaster during early metamorphosis were examined by assaying whole animals at stages that span two pulses of ecdysone. Microarrays were constructed containing 6240 elements that included more than 4500 unique cDNA expressed sequence tag (EST) clones along with a number of ecdysone-regulated control genes having predictable expression patterns. These ESTs represent approximately 30% to 40% of the total estimated number of genes in the Drosophila genome. In order to gauge expression levels, microarrays were hybridized with fluorescent probes derived from polyA+ RNA isolated from developmentally staged animals. The time points examined are relative to PF, which last approximately 15 to 30 min, during which time the larvae cease to move and evert their anterior spiracles. Nineteen arrays were examined representing six time points relative to PF: one time point before the late larval ecdysone pulse; one time point just after the initiation of this pulse (4 hours BPF), and time points at 3, 6, 9, and 12 hours after PF (APF). The prepupal pulse of ecdysone occurs 9 to 12 hours APF (White, 1999).

In order to manage, analyze, and disseminate the large amount of data, a searchable database was constructed that includes the average expression differential at each time point. The analysis set consists of all elements that reproducibly fluctuate in expression threefold or more at any time point relative to PF, leaving 534 elements containing sequences represented by 465 ESTs and control genes. More than 10% of the genes represented by the ESTs display threefold or more differential expression during early metamorphosis. This may be a conservative estimate of the percentage of Drosophila genes that change in expression level during early metamorphosis, because of the stringent criteria used for their selection (White, 1999).

To interpret these data, genes were grouped according to similarity of expression patterns by two methods. The first relied on pairwise correlation statistics, and the second relied on the use of self-organizing maps (SOMs). Differentially expressed genes fall into two main categories. The first category contains genes that are expressed at >18 hours BFP (before the late larval ecdysone pulse) but then fall to low or undetectable levels during this pulse. These genes are potentially repressed by ecdysone and make up 44% of the 465 ESTs identified in this set. The second category consists of genes expressed at low or undetectable levels before the late larval ecdysone pulse but then are induced during this pulse. These genes are potentially induced by ecdysone and make up 31% of the 465 ESTs. Consequently, 75% of genes that changed in expression by threefold or more do so during the late larval ecdysone pulse that marks the initial transition from larva to prepupa. This result is consistent with the extreme morphological changes that are about to occur in these animals. There are clearly discrete subdivisions of gene expression within these categories (White, 1999).

Gene expression changes during metamorphosis also foreshadow both larval muscle breakdown and adult myogenesis. At approximately 2 hours APF, the anterior larval musculature begins to break down. This breakdown lasts until approximately 6 hours APF. Genes encoding both structural and regulatory components of muscle formation are down-regulated as early as 4 hours BPF (see Muscle-specific genes regulated during metamorphosis). In addition to the repression of genes encoding components of thin and thick filaments, genes encoding other muscle-specific molecules are also repressed, including factors that compose the mesh in which these filaments lie and regulatory factors involved in the specification of muscle tissue. The mRNAs of all these repressed genes decrease substantially many hours before histolysis of the anterior larval muscles and therefore predict the occurrence of this morphological event well before it begins. Twenty-four hours APF, adult myogenesis is well underway. The genes DMef-2, bagpipe, and tinman are all up-regulated at 12 hours APF from the baseline at PF, coincident with the prepupal pulse of ecdysone. It is suggested that induction of these regulatory factors initiates the development of the adult musculature, which will establish itself several hours later (White, 1999).

Mef-2 in cultured cells

Schneider SL2 cells activate the myogenic program in response to the ectopic expression of daughterless alone, as indicated by exit from the cell cycle, syncytia formation, and the presence of muscle myosin fibrils. Myogenic conversion can be potentiated by the coexpression of Drosophila Mef2 and nautilus with daughterless. In RT-PCR assays Schneider cells express two mesodermal markers, Nautilus and Mef2 mRNAs, as well as very low levels of Daughterless mRNA but no Twist. Full-length RT-PCR products for Nautilus and Mef2 encode immunoprecipitable proteins. RNA-i was used to demonstrate that both endogenous nautilus expression and Mef2 expression are required for the myogenic conversion of Schneider cells by daughterless. Coexpression of twist blocks conversion by daughterless but twist dsRNA has no effect. These results indicate that Schneider cells are of mesodermal origin and that myogenic conversion with ectopic expression of daughterless occurs by raising the levels of Daughterless protein sufficiently to allow the formation of Nautilus/Daughterless heterodimers. The effectiveness of RNA-i is dependent upon protein half-life. Genes encoding proteins with relatively short half-lives (10 h), such as Nautilus or Hsf, are efficiently silenced, whereas more stable proteins, such as cytoplasmic actin or beta-galactosidase, are less amenable to the application of RNA-i. These results support the conclusion that Nautilus is a myogenic factor in Drosophila tissue culture cells with a functional role similar to that of vertebrate MyoD. This is discussed with regard to the in vivo functions of Nautilus (Wei, 2000).

Effects of Mutation or Deletion

Complete loss of MEF2 protein in homozygous mutant embryos results in a dramatic absence of myosin heavy chain (MHC)-expressing myoblasts and lack of differentiated muscle fibers. Examination of earlier events involved in muscle development indicates that the specification and early differentiation of somatic muscle precursors are not affected because even-skipped-, nautilus-, and beta 3-tubulin-expressing myoblasts are present. However, these partially differentiated cells are unable to undergo further differentiation to form muscle fibers in the absence of Mef2.

The later aspects of differentiation of the visceral mesoderm and the heart are also disrupted in Mef2 mutant embryos, although the specification and early development of these tissues appear unaffected. Midgut morphogenesis is disrupted in the mutant embryos, presumably as a consequence of abnormal development of the visceral mesoderm. In the heart, the formation of cardial cells is not affected by the absence of Mef2, but differentiation here is deficient. even-skipped expressing paracardial cells are present in Mef2 mutants.

Mef2 deficiency is reminiscent of myogenin phenotype in mice. Myogenin is needed for the terminal differentiation of myoblasts, but not for their determination. Roles in myoblast determination have been ascribed to MyoD and Myf-5 (Bour, 1995).

Embryos deficient for MEF2 protein, due to a deletion of upstream transcriptional control sequences, fail to form muscle, suggesting that the gene is required for muscle cell differentation. In the somatic muscle lineage, Mef2 is required for the formation and patterning of body wall muscle. In the absence of somatic myogenesis, there is extensive apoptosis among the myoblast cell population. In contrast, in the cardiac muscle lineage, morphogenesis of the dorsal vessel occurs normally, but the three myosin subunit genes are not expressed. MeF2 is also expressed in adepithelial cells. Rare Mef2 transheterozygous mutant adults fail to fly, consistent with defects observed in the indirect flight muscles (Ranganayakulu, 1995).

The expression of the MyoD gene homolog, nautilus (nau), in the Drosophila embryo defines a subset of mesodermal cells known as the muscle 'pioneer' or 'founder' cells. These cells are thought to establish the future muscle pattern in each hemisegment. Founders appear to recruit fusion-competent mesodermal cells to establish a particular muscle fiber type. In support of this concept every somatic muscle in the embryo is associated with one or more nautilus-positive cells. However, because of the lack of known (isolated) nautilus mutations, no direct test of the founder cell hypothesis has been possible. Toxin ablation and genetic interference by double-stranded RNA (RNA interference or RNA-i) have been used to determine both the role of the nautilus-expressing cells and the nautilus gene, respectively, in embryonic muscle formation. In the absence of nautilus-expressing cells muscle formation is severely disrupted or absent. A similar phenotype is observed with the elimination of the nautilus gene product by genetic interference upon injection of nautilus double-stranded RNA (Misquitta, 1999).

The results from the injection of nautilus dsRNA point to a more general approach for the analysis of gene function during Drosophila development and suggest that the RNA interference method essentially would mimic a gene knock-out in the injected generation of Drosophila embryos. To test this idea a variety of cDNA clones were obtained representing a maternal gene expressed in the embryo (daughterless); additional genes involved in myogenesis (S59, DMEF2); homeobox genes (engrailed and S59); a gene important for gastrulation (twist), and a gene expressed in the adult eye (white). This panel of genes covers most stages of Drosophila development. twist was initiatially tested because the mutant has a clear phenotype that is easy to score when compared with wild-type larva. The injection of twist dsRNA (the complete coding region) into embryos produces a twisted larval phenotype that is indistinguishable from the original twist mutation. Similarly, injection of the first 1,200 bp of engrailed dsRNA produces the compressed dentical belt pattern characteristic of an engrailed null mutant. Daughterless mRNA is both maternally loaded and expressed zygotically, and the mutant phenotype produces very characteristic disruptions in the central nervous system (CNS) and peripheral nervous system (PNS). It has been shown previously that mex3, a maternally loaded RNA in C. elegans, can be ablated by dsRNA injection into the gonads. daughterless dsRNA (complete coding region) was injected and the characteristic neuronal phenotypes were sought by using the mAb MAB 22C10. The CNS as well as the PNS were disrupted to varying degrees in the injected embryos. The severity of the phenotype consistently shows a CNS disruption with a variable PNS pattern, possibly reflecting the fact that the CNS is formed before the PNS. This result suggests that maternally loaded as well as zygotically expressed RNA can be affected by RNA-i in Drosophila. The homeobox gene S59 marks a subset of muscle founder cells for 5 of 29 muscles in each hemisegment of the embryo corresponding to muscles 5, 18, 25, 26, and 27. Embryos with an S59 lacZ transgene marking muscles 18 and 25 were injected with S59 dsRNA (complete coding region). In this case, the S59-specific lacZ antibody-staining pattern is abolished. The total muscle pattern for embryos injected with S59 dsRNA, although disrupted, still shows the presence of poorly organized muscle groups in each hemisegment. This is unlike the almost complete absence of muscle observed with the injection of nautilus dsRNA. DMEF2, a member of the MADS domain transcription factor family, is essential for muscle formation in Drosophila. The DMEF2 / embryo has no muscle and is missing the characteristic gut constrictions found in the uninjected embryo. Injection of DMEF2 dsRNA (complete coding region) results in embryos that lack any detectable muscle and an absence of gut morphology (Misquitta, 1999).

mef2 genes encode alternatively spliced transcription factor isoforms that function in muscle differentiation in both Drosophila and vertebrates. Drosophila mef2 (Dmef2) has been shown to be required for the differentiation of a variety of distinct muscle types. However, many possible aspects of its function in muscle remain unexplored. There has also been no analysis in vivo of the activity of different Mef2 isoforms in any species. This investigation centers on the role of different levels of DMef2 in regulating diverse events of muscle differentiation in the Drosophila embryo and on the functional significance of Dmef2 alternative splicing. The GAL4/UAS system was used to both misexpress and overexpress individual DMef2 isoforms and to rescue the different aspects of the Dmef2 mutant phenotype. Ectopic ectodermal expression of DMef2 activates muscle gene expression and inhibits epidermal differentiation. Overexpression of DMef2 in the mesoderm disrupts differentiation of the somatic and visceral muscle and the heart. The use of different DMef2 levels in rescue experiments reveals an activity range compatible with differentiation of the different muscle types: the consequence of too little or too much DMef2 activity is disrupted differentiation. These rescue experiments also reveal that distinct DMEF2 thresholds are required for different properties within a cell and also for different cells within a muscle type and for different muscle types. For example, relatively low levels of DMef2 rescue myosin expression in the visceral muscle, together with, in almost all embryos, the midgut constrictions. Higher levels are required for complete rescue of the alary muscles, while the pharyngeal muscle and somatic musculature are only fully rescued with the higher levels produced at 29 degrees C. Complete rescue of myosin expression in the heart is only achieved in about 35% of the embryos at 29 degrees C., indicating that this tissue is the most difficult to rescue. Finally, each isoform functions equivalently in these experiments, including in the stringent test of rescue of the Dmef2 mutant phenotype (Gunthorpe, 1999).

The Him gene inhibits the development of Drosophila flight muscles during metamorphosis

During Drosophila metamorphosis some larval tissues escape the general histolysis and are remodelled to form adult tissues. One example is the dorso-longitudinal muscles (DLMs) of the indirect flight musculature. They are formed by an intriguing process in which residual larval oblique muscles (LOMs) split and fuse with imaginal myoblasts associated with the wing disc. These myoblasts arise in the embryo, but remain undifferentiated throughout embryogenesis and larval life, and thus share characteristics with mammalian satellite cells. However, the mechanisms that maintain the Drosophila myoblasts in an undifferentiated state until needed for LOM remodelling are not understood. This study shows that the Him gene is expressed in these myoblasts, but is undetectable in developing DLM fibers. Consistent with this, it was found that Him could inhibit DLM development: it inhibited LOM splitting and resulted in fiber degeneration. Then a balance between mef2, a positive factor required for proper DLM development, and the inhibitory action of Him, was uncovered. Mef2 suppressed the inhibitory effect of Him on DLM development, while Him could suppress the premature myosin expression induced by mef2 in myoblasts. Furthermore, either decreased Him function or increased mef2 function disrupted DLM development. These findings, together with the co-expression of Him and Mef2 in myoblasts, indicate that Him may antagonise mef2 function during normal DLM development and that Him participates in a balance of signals that controls adult myoblast differentiation and remodelling of these muscle fibers. Lastly, evidence is provided for a link between Notch function and Him and mef2 in this balance (Solera, 2009).

This paper analysed events in the development of the DLMs, components of the indirect flight musculature, during Drosophila adult myogenesis. Him was found to be expressed in myoblasts, but not in developing adult muscle fibers, and consistent with this pattern of expression it was found that Him could inhibit DLM development. Template splitting was inhibited and subsequently remaining muscle fibers degenerated. This phenotype resembles that found when the myoblasts are removed by a genetic manipulation. It is inferred from this phenotypic similarity that over-expression of Him makes these myoblasts functionally defective. It was also found that this phenotype due to Him is suppressed by mef2. This genetic interaction, together with previous observations that mef2 hypomorph mutants also have impaired LOM splitting, is consistent with Him antagonising Mef2 activity in these myoblasts. It is noted, however, that in contrast to mef2 hypomorphic mutants, Him can induce degeneration of the DLMs. This could be because 1151-Gal4 driving UAS-Him results in a lower level of Mef2 activity than in the hypomorphic mef2 mutants, and/or because Him induces DLM degeneration by another route. The rescue of the degeneration phenotype, as well as of template splitting, by mef2 suggests that the effect of Him is at least substantially through the mef2 pathway (Solera, 2009).

It is apparent that there must be a mechanism to inhibit Mef2 function in the committed, but undifferentiated adult myoblasts associated with the wing discs, but it was not known how this is achieved. The current findings suggest that Him could be such an inhibitor. Him could antagonise the mef2-induced expression of myosin, a classic marker of muscle differentiation, in disc-associated myoblasts. Moreover, in normal development, Him is already expressed in the myoblasts when Mef2 is first expressed in the third larval instar. Him and Mef2 also remain co-expressed in the myoblasts during the early phase of pupal life, but later, as the muscle fibers differentiate, Mef2 is expressed in the fibers whereas Him is not, although it continues in surrounding myoblasts. This arrangement where Mef2 is expressed together with an inhibitor suggests a capacitor-like situation in which there could be a burst of Mef2 activity, a discharge, when the inhibition is relieved. This could result in a co-ordinated onset of a phase of the muscle differentiation programme (Solera, 2009).

This study has revealed a balance of Him and mef2 activities in the control of adult myoblast differentiation. Thus, additional Him expression, like reduced mef2 function, leads to disrupted DLM development, and the effect of Him can be suppressed by mef2. Moreover, although mef2 has a positive role in adult myogenesis, it is required for proper DLM development, it was found that over-expression of mef2 affected DLM formation and the total number of DLM fibers was somewhat reduced compared to wild type. It was also found that reduced expression of the Mef2 inhibitor Him had a similar effect. This indicates that too much of a positive signal, or too little of a negative, can affect muscle development. Taken together with other results, this suggests that the balance of antagonistic activities, which includes mef2 and Him, has to be precisely controlled to ensure correct implementation of the differentiation programme (Solera, 2009).

One other known input into the control of adult myogenesis is Notch. It has been shown to inhibit adult muscle differentiation, and it has been suggested that it may do this through twist, although much remains unclear about the molecular pathway(s) involved. The results link Notch with two other regulators. It was found that mef2 over-expression could partially suppress the inhibitory effect of N-intra on DLM development, and that N-intra resulted in Him expression in muscle fibers. The latter might be direct regulation of gene expression because a region upstream of the Him gene contains several binding sites for Su(H), a nuclear effector of Notch signalling, and either deletion of this region or a Su(H) genetic background results in down-regulation of Him expression in wing discs. Together these results indicate that Notch can upregulate Him expression, which in turn can inhibit mef2 function, and this will result in inhibition of DLM development (Solera, 2009).

These findings further understanding of the factors that influence the programme of differentiation during adult Drosophila myogenesis. They also pave the way for future studies into the mechanisms that control this intriguing system of muscle development, which includes tissue remodelling and is effectively a type of regeneration, and which also has certain parallels with mammalian muscle post-natal growth and repair. This work also illustrates the significance of a balance of activities in how adult progenitor cells might be maintained in a committed, but undifferentiated state and then be triggered to enter the differentiation pathway. This is of broad significance both in development and in potential applications of stem cell biology (Solera, 2009).

Differential requirements for Myocyte Enhancer Factor-2 during adult myogenesis in Drosophila

Identifying the genetic program that leads to formation of functionally and morphologically distinct muscle fibers is one of the major challenges in developmental biology. In Drosophila, the Myocyte Enhancer Factor-2 (MEF2) transcription factor is important for all types of embryonic muscle differentiation. This study investigated the role of MEF2 at different stages of adult skeletal muscle formation, where a diverse group of specialized muscles arises. Through stage- and tissue-specific expression of Mef2 RNAi constructs, it was demonstrated that MEF2 is critical at the early stages of adult myoblast fusion: mutant myoblasts are attracted normally to their founder cell targets, but are unable to fuse to form myotubes. Interestingly, ablation of Mef2 expression at later stages of development showed MEF2 to be more dispensable for structural gene expression: after myoblast fusion, Mef2 knockdown did not interrupt expression of major structural gene transcripts, and myofibrils were formed. However, the MEF2-depleted fibers showed impaired integrity and a lack of fibrillar organization. When Mef2 RNAi was induced in muscles following eclosion, no adverse effects of attenuating Mef2 function were found. It is concluded that in the context of adult myogenesis, MEF2 remains an essential factor, participating in control of myoblast fusion, and myofibrillogenesis in developing myotubes. However, MEF2 does not show a major requirement in the maintenance of muscle structural gene expression. findings point to the importance of a diversity of regulatory factors that are required for the formation and function of the distinct muscle fibers found in animals (Bryantsev, 2012).

This study demonstrates an important requirement for MEF2 in the formation of the adult muscles, where early reduction in MEF2 levels blocks myoblast fusion, and obliterates formation of the adult somatic muscles. These findings are more severe than had been previously reported for a Mef2 temperature-sensitive combination; why should this be the case? The most reasonable explanation for this discrepancy is that the temperature-sensitive mutants, while severely impacting MEF2 function, nevertheless still showed some low level of MEF2 activity and myogenic potential, and that this low level of MEF2 function was sufficient to support adult myogenesis. An additional factor to consider is the period of time that it takes for the first steps of myogenesis to occur, at the adult stage versus the embryonic stage. In embryos, the fusion process is completed within a time window of just a few hours; whereas in adult myogenesis, fusion proceeds over the course of at least 12 h of pupal development. It is feasible then, that low levels of sustained MEF2 function over an extended period of time are sufficient to support the earliest steps of adult muscle development, while such an effect is not possible in embryonic myogenesis, due to its strictly limited timing (Bryantsev, 2012).

Data is presented indicating that there is yet some residual MEF2 accumulation in the skeletal muscles of some of the RNAi lines that were characterized, since haploinsufficiency for Mef2, or use of a double-RNAi, exacerbated the myofibrillar defects in post-fusion Mef2 knockdowns. Nevertheless, these levels must be extremely low, given the immunofluorescent data indicating that indeed levels of MEF2 are reduced to background detection levels from as early as 24 h APF time-points. More importantly, neither haploinsufficiency for Mef2 nor the double-RNAi background reduced the levels of Act88F and Mhc expression. It is therefore concluded that there are clear stage-specific requirements for this important transcription factor, where high levels of MEF2 are required for early myogenesis, but later myogenesis appears to proceed with only minimal levels (if any) of MEF2 function required (Bryantsev, 2012).

Fusion defects occur in null Mef2 mutants during embryogenesis, however such a phenotype has not yet been reported for adult myogenesis. This study, demonstrating a requirement for MEF2 in adult myoblast fusion, therefore underlines a similarity between these two distinct phases of Drosophila muscle development. The data suggest that the abolition of myoblast fusion is at least partly due to the significant down-regulation of singles bar (sing) transcripts, in samples of purified myoblasts plus myotubes isolated from cryosections. Sing expression was also decreased in Mef2 mutant embryos (Sandmann, 2006), conferring another level of similarity between the molecular mechanisms regulating myoblast fusion in embryonic and adult myogenesis. Importantly, the expression levels of two other tested fusion genes, blow and mbc, did not show any detectable alterations in expression in response to Mef2 knockdown. It is concluded that MEF2 is not a key regulator for multiple fusion genes, although it certainly is essential for fusion via activation of sing transcription. It is noted that mbc is not exclusively expressed in developing muscles, and is in fact more broadly-distributed, reducing the likelihood that its expression solely responds to MEF2 activity. Whether there are more MEF2-dependent fusion genes, and whether MEF2 is a direct transcriptional activator of such genes, is yet to be fully established at either the embryonic or the adult stages (Bryantsev, 2012).

The findings also demonstrate that the transcriptional initiation of genes encoding adult myofibrillar protein genes depends upon MEF2 being present early in muscle formation. Whereas a sample from control muscle fusion templates plus myoblasts showed robust expression of both Act88F and Mhc at 30 h APF, Mef2-knockdown samples showed sparse activation of these muscle structural genes. The findings are consistent with the role of Mef2 in embryonic myogenesis, where the majority of muscle-specific genes fail to become expressed in the absence of MEF2 function. Clearly, a role for MEF2 in the early stages of muscle formation appears to be a commonality between embryonic and adult early myogenesis. Nevertheless, some intermediate factors must also be important in this initiation process, since it has yet to be demonstrated that adult actin gene expression is directly responsive to MEF2 (Bryantsev, 2012).

In agreement with earlier data, there is clearly a more moderate requirement for MEF2 function in formation of the adult muscles following myoblast fusion. Despite MEF2 being immunologically undetectable shortly after fusion in RNAi animals, significant myogenesis occurs, and this process results in the formation of muscles in pharate adults. This finding is in contrast to Mef2 knockdown during the early stages of adult myogenesis, where it was absolutely required for activation of fibrillogenesis (Bryantsev, 2012).

Importantly, adult-specific actin gene expression does not become stalled or aborted in the Mef2 knockdown animals, as can be judged based on three major observations: (1) F-actin clearly accumulates in Mef2 knockdown myofibers; (2) the tissue-specific drivers, derived from adult muscle actin genes, function effectively to suppress MEF2 accumulation until the end of myogenesis, and therefore are not dependent on MEF2 activity; and (3) quantitative RT-PCR analysis did not reveal statistically significant differences in Act88F expression levels between experimental and control samples at late time-points of myogenesis (72 h and 90 h APF), even in a Mef2 haploinsufficient background and in a double-RNAi background. These data provide further support for the notion that MEF2 is at most minimally required for the sustained expression of some adult muscle genes. Moreover, neither the Act88F nor the Act79B enhancers contain canonical MEF2 binding sites, and neither enhancer is activated by MEF2 in transient co-transfection assays (Bryantsev, 2012).

In the light of the demonstrated independence of adult actin gene transcription from Mef2 expression, one must ask how known MEF2-dependent genes, with general muscle expression, respond to Mef2 silencing during adult myogenesis? In this study Mhc was used as an example of such a pan-muscle-specific gene, since in the embryo initiation of Mhc expression depends on Mef2 activity. The data, however, indicate that MEF2 depletion does not alter Mhc expression at statistically significant levels, even under exacerbated conditions. It is concluded that while MEF2 is required to initiate expression of Mhc, MEF2 is likely to be just one of the factors participating in the maintenance of Mhc expression (Bryantsev, 2012).

The results are supported by the studies of others, in which the Mhc gene was shown to receive regulatory input from numerous evolutionarily conserved cis-regulatory elements scattered throughout its complex regulatory region, expanding from as far as 10-kb upstream of the transcription start site and into the first intron. Moreover, some of the cloned Mhc enhancer elements, lacking MEF2 sites, directed myofiber-specific reporter gene expression in adult flies. Apparently, maintenance of Mhc expression in adults depends on multiple transcription factors (Bryantsev, 2012).

Besides Mhc, other pan-specifically expressed muscle genes have been investigated for transcriptional regulatory elements. Tropomyosin-1 and -2, Troponin T, and Paramyosin/mini-Paramyosin also possess complex enhancers comprising multiple regulatory elements that can be activated in a myofibril-specific fashion, often independently of MEF2 sites, suggest that MEF2, while being important for activation of muscle structural genes, becomes less critical in their maintenance, which concurs with the current observations. The latter postulate can explain why enhancers containing generic MEF2 sites, being functionally active in cell culture co-transfection assays, fail to support muscle-specific expression of the LacZ reporter gene in transgenic animals; and why a single MEF2 site with flanking sequences, taken out of the context of a functional embryonic Act57B enhancer, is not capable of driving reporter gene expression in embryonic muscles (Bryantsev, 2012).

Nevertheless, MEF2 remains a key regulator of early steps of myogenesis in Drosophila. According to the experimental data, MEF2 functions in early stages of adult myogenesis to support myoblast fusion, and it is postulated that at this time MEF2 also begins to activate additional muscle-specific regulatory factors that then go on to activate muscle structural genes during the remainder of adult myogenesis (Bryantsev, 2012).

Very few factors, other than MEF2, directly regulate muscle structural genes in Drosophila. Chorion Factor-2 (CF2) is an important MEF2 co-factor for muscle gene expression in the embryo/larva, and it does play a role in adult myogenesis as well. The Cf2 gene is genetically downstream of Mef2, while Pdp1 regulation is complicated due to the presence of multiple alternative promoters, but late accumulation of products of this gene in skeletal muscles during embryogenesis may suggest that it is also MEF2-dependent. Thus, MEF2 can be an initiator for expression of additional myogenic regulators, and these factors can later acquire their own positive auto-regulatory loop. In adults, the transcriptional co-activator vestigial, participating in IFM development, is activated by MEF2 in adult-specific myoblasts and then participates in self-activation. It is plausible that other transcriptional regulators participating in myofiber-specific myogenesis first receive activation of their expression from MEF2 — to be later maintained by different means (Bryantsev, 2012).

Further, there is clearly a subset of muscle protein genes that are activated after myoblast fusion and that must nevertheless be significantly affected by loss of MEF2 function. Evidence for this comes from the pathology that does occur in Mef2-knockdown myofibers, which includes loss of proper myofibril organization, detachment of muscle fibers from the attachment sites, and separation of neighboring myofibers. Some of the Mef2-knockdown phenotypes were clearly visible even after 18 h of Mef2 RNAi initiation, indicating that there must be targets that are heavily dependent on MEF2 activity. Identification of such target genes should become a focus of future research efforts (Bryantsev, 2012).

In the animals in which Mef2 knockdown was induced post eclosion, the severe changes in lifespan conferred by the IR 5039 are not considered as being a true phenotype of MEF2 loss. Most likely, this phenotype is a reflection of RNAi off-target activity. Thus, post-eclosion, there appears to be relatively little requirement for MEF2, at least over the time period and under the environmental conditions that were tested. This finding is consistent with the classical observations that demonstrated that there is extremely low turnover of thoracic proteins following eclosion, and this study additionally demonstrated that sustained expression of Act88F is not required following the first day of adult life. It cannot be ruled out that MEF2 may be participating in other aspects of muscle physiology, besides structural protein turn-over. Hence, MEF2 might be involved in expression of muscle-specific enzymes, controlling muscle metabolism. Another intriguing possibility could be a potential participation of MEF2 in muscle repair after an extensive muscle exercise such as long-term flying, or pathology. The sustained presence of MEF2 in mature adult muscles argues strongly for a role in these cells, which additional studies should be designed to uncover (Bryantsev, 2012).

Vertebrate slow/fast twitch muscles consist of fibers with different functional, metabolic, and molecular properties, just as Drosophila adult muscles are fine-tuned in order to carry out their unique physiological functions. In both cases, it is still unclear how initial fiber types are specified in the nascent muscle. In mammals, there is evidence that fiber-type switching relies on MEF2 working in combination with other transcription factors to selectively activate muscle structural genes. Given the broad conservation in the basic mechanisms of myogenesis, as exemplified by the role of MEF2 in muscle differentiation from flies to vertebrates, it is anticipated that studying adult myogenesis in Drosophila, which had been hindered by the lack of appropriate molecular tools, now can start providing new genetic data on myofiber-specific development (Bryantsev, 2012).

MEF2 is an in vivo immune-metabolic switch

Infections disturb metabolic homeostasis in many contexts, but the underlying connections are not completely understood. To address this, paired genetic and computational screens were used in Drosophila to identify transcriptional regulators of immunity and pathology and their associated target genes and physiologies. It was shown that Mef2 is required in the fat body for anabolic function and the immune response. Using genetic and biochemical approaches, it was found that MEF2 is phosphorylated at a conserved site in healthy flies and promotes expression of lipogenic and glycogenic enzymes. Upon infection, this phosphorylation is lost, and the activity of MEF2 changes-MEF2 now associates with the TATA binding protein to bind a distinct TATA box sequence and promote antimicrobial peptide expression. The loss of phosphorylated MEF2 contributes to loss of anabolic enzyme expression in Gram-negative bacterial infection. MEF2 is thus a critical transcriptional switch in the adult fat body between metabolism and immunity (Clark, 2013).

This study identified Mef2 as a factor critical for energy storage and the inducible immune response in the Drosophila fat body. Many infection-induced antimicrobial peptides depend on Mef2 for normal expression. In consequence, flies lacking Mef2 activity in the fat body are severely immunocompromised against a variety of infections. Mef2 sites are also associated with genes encoding key enzymes of anabolism, and Mef2 is required for normal expression of these genes; consequently, flies lacking Mef2 function in the fat body exhibit striking reductions in the total levels of triglyceride and glycogen. These two groups of target genes are counterregulated during infection; the anabolic targets of Mef2 are reduced in expression when antimicrobial peptides are induced. Fat body MEF2 was shown to exist in two states with distinct physiological activities. In uninfected animals, MEF2 is phosphorylated at T20 and can promote the expression of its metabolic targets. In infected animals, T20 is dephosphorylated, and MEF2 associates with the TATA-binding protein to bind a compound MEF2-TATA sequence found in the core promoters of antimicrobial peptides. The loss of T20-phosphorylated MEF2 promotes the loss of anabolic transcripts in flies with Gram-negative bacterial infection. These data, taken together, suggest that the central role of MEF2 in promoting fat body anabolism and immune activity reflects a switch between distinct transcriptional states regulated, at least in part, by differential affinity for TBP determined by T20 phosphorylation (Clark, 2013).

The signaling mechanisms regulating T20 phosphorylation and MEF2-TBP association are clearly of critical importance. The ability of p70 S6K to phosphorylate this residue is congruent with the ability of S6K to enhance anabolism and repress catab- olism in response to nutrient signals (Laplante, 2012). However, others have shown T20 phosphorylation by PKA, suggesting that T20 phosphorylation may be regulated by more than one pathway in vivo. The role of TAK1 may be similarly complex. TAK1 is required for formation of the MEF2-TBP complex upon Gram-negative infection, but this effect may be indirect. For example, reduced S6K phosphorylation after infection may result from insulin resistance driven by TAK1 via JNK. TAK1- dependent JNK activation is required for normal AMP induction in vivo, but it remains possible that some novel pathway is the critical connection between TAK1 and MEF2-TBP complex formation (Clark, 2013).

In mammals, in addition to hematopoietic roles, Mef2c regulates B cell proliferation upon antigen stimulation, and Mef2d regulates IL2 and IL10 in T cells. The possibility that Mef2 family proteins might be important direct activators of innate responses has not previously been examined. This study shows that Mef2 is a core transcriptional component of the innate immune response of the adult fly. Equally, vertebrate Mef2 family proteins are critical regulators of muscle metabolism, activated by physical activity to promote expression of PGC-1a and the glucose transporter Glut4. Glut4 regulation by MEF2 is known in adipose tissue as well as in muscle 1998); it is an intriguing possibility that MEF2 is as important a regulator of adipose metabolism in vertebrates as it has been show to be in flies (Clark, 2013).

Infection-induced metabolic disruption leading to cachexia is present in vertebrates as well as in insects, most notoriously in Gram-negative sepsis and persistent bacterial infections such as tuberculosis. The current data suggest that wasting seen after infection may be due, in part, to the requirement for MEF2 to serve different transcriptional functions in different conditions; the MEF2 immune-metabolic transcriptional switch may be a mechanistic constraint that forces the fly into metabolic pathophysiology in contexts of persistent immune activation. Alternatively, the loss of MEF2-driven anabolic transcripts due to infection may be productive, either by altering systemic energy usage or by increasing the production or release of one or more antimicrobial metabolites. Recent work has highlighted a distinction between 'resistance' type immune mechanisms, in which the host attempts to eradicate an invading organism, and 'tolerance' type mechanisms, in which the host response is oriented toward reducing the damage done by infection. The distinct metabolic and immune requirements for MEF2, combined with the obligation on the part of the host to raise some measure of resistance to systemic infection, may limit the achievable level of tolerance in persistent infections (Clark, 2013).

A rapid one-generation genetic screen in a Drosophila model to capture rhabdomyosarcoma effectors and therapeutic targets

Rhabdomyosarcoma (RMS - see Drosophila as a Model for Human Diseases: Rhabdomyosarcoma) is an aggressive childhood malignancy of neoplastic muscle-lineage precursors that fail to terminally differentiate into syncytial muscle. The most aggressive form of RMS, Alveolar-RMS (A-RMS), is driven by misexpression of the PAX-FOXO1 oncoprotein, which is generated by recurrent chromosomal translocations that fuse either the PAX3 or PAX7 gene (homologs of Drosophila Paired) to FOXO1 (homolog of Drosophila Foxo). The molecular underpinnings of PAX-FOXO1-mediated RMS pathogenesis remain unclear, however, and clinical outcomes poor. This study reports a new approach to dissect RMS, exploiting a highly efficient Drosophila PAX7-FOXO1 model uniquely configured to uncover PAX-FOXO1 RMS genetic effectors in only one generation. With this system, a comprehensive deletion screen was performed against the Drosophila autosomes, and mutation of Mef2, a myogenesis lynchpin in both flies and mammals, was demonstrated to dominantly suppresses PAX7-FOXO1 pathogenicity and act as a PAX7-FOXO1 gene target. Additionally, mutation of mastermind, a gene encoding a MEF2 transcriptional co-activator, was shown to similarly suppress PAX7-FOXO1, further pointing towards MEF2 transcriptional activity as a PAX-FOXO1 underpinning. These studies show the utility of the PAX-FOXO1 Drosophila system as a robust one-generation (F1) RMS gene discovery platform and demonstrate how Drosophila transgenic conditional expression models can be configured for the rapid dissection of human disease (Galindo, 2014: PubMed).

Given the critical role that the PAX-FOXO1 fusion oncoprotein plays in RMS, this study focuses on PAX-FOXO1 as an entry-point for designing a transgenic Drosophila RMS-related model that would be amenable to forward genetic screening and RMS gene discovery. To bypass the issue of cumbersome multigenerational screening schemes that would normally be required, a Gal80 X-linked chromosomal transgene was incorporated to generate a viable screening Gal4/UAS-PAX-FOXO1 master stock that allows for the rapid identification of PAX-FOXO1 genetic modifiers in a single genetic cross (Galindo, 2014).

With this platform, new PAX-FOXO1 pathogenesis underpinnings were probed. Though very similar in molecular structure, PAX3-FOXO1− and PAX7-FOXO1−positive RMS demonstrate differing clinical behaviors, as PAX3-FOXO1 tumors are more common and notoriously aggressive. Consequently, PAX3-FOXO1 is the PAX-FOXO1 fusion most commonly investigated in vertebrate models. This study focuses on PAX7-FOXO1 in the Drosophila system, which demonstrates phenotypes that are better penetrant and experimentally tractable due to the fact that human PAX7 demonstrates slightly greater sequence identity to fly PAX3/7 than does human PAX3. Additionally, as no other animal models of PAX7-FOXO1 presently exist, the fly PAX7-FOXO1 model also conveniently serves as a complement to vertebrate PAX3-FOXO1 models (Galindo, 2014).

The extent to which observations from the PAX7-FOXO1 fly model would impact the clinically more aggressive PAX3-FOXO1 RMS subtype, as well as PAX-FOXO1-negative (embryonal) RMS, is unknown. Notably, previous studies show that genetic modifiers identified from the Drosophila system impact PAX3-FOXO1 RMS oncogenesis and tumorigenesis. Furthermore, unpublished studies suggest that fly PAX7-FOXO1 genetic modifiers are similarly involved in Embryonal RMS. These findings provide marked validation for the applicability and value of this genetic fly system to human RMS (Galindo, 2014).

Interestingly, though PAX7-FOXO1 induces expression of the late myogenic differentiation marker MHC, PAX-FOXO1 RMS myoblasts in culture and in vivo demonstrate only partial differentiation with little-to-no MHC expression. In considering this discrepancy, it should be first noted that PAX-FOXO1 is a relatively weak driver of RMS in culture and in vivo and requires additional/sequential genetic aberrations to induce oncogenic transformation. Thus, secondary mutations might be necessary to force the strength of RMS myoblast differentiation-arrest seen in human RMS tumors; by contrast, the PAX7-FOXO1 model of this study differs in that the system is free of any additional background mutations. Second, earlier studies show that expression of PAX3-FOXO1 in mouse embryonic cultured cells induces the formation of MHC-positive myocytes and myotube, similar to the Drosophila system in this study as the da-Gal4/UAS-PAX7-FOXO1 expression system targets undifferentiated embryonic primordia. Uncovering of the genetic/molecular sequence of RMS pathogenesis and the cell(s) origin will shed further insight into the underlying mechanisms that account for the myoblast differentiation arrest phenotypes seen in RMS in vivo (Galindo, 2014).

The differentiation and fusion of myoblasts into postmitotic, syncytial muscle requires that the bHLH myogenic regulatory factors (MRFs: Myf5, Mrf4, MyoD, and Myogenin) interact with E-proteins, which drive and regulate critical aspects of myogenic fate determination. The MRFs subsequently interact with the MEF2 transcription factors that, although lacking intrinsic myogenic activity, cooperate with the MRFs to synergistically activate muscle-specific genes and the downstream myogenic terminal differentiation program. Vertebrates possess four MEF2 family member genes (-A, -B, -C, -D), which demonstrate complex overlapping spatial and temporal expression patterns in embryonic and adult tissues, with greatest expression levels seen in striated muscle and brain. Because of genetic redundancy and overlapping expression patterns of the MEF2 genes, interrogating individual MEF2 gene activity in mammals is experimentally challenging, with loss-of-function mutation studies revealing only limited insights into MEF2 gene function in tissues in which the MEF2 genes do not overlap/compensate. Conveniently, flies possess only one Mef2 gene (D-Mef2) and serve as an excellent model system to delineate MEF2’s critical role in myogenesis. The study speculates that the lack of Mef2 redundancy in flies provides a marked experimental advantage in isolating D-Mef2 as a PAX7-FOXO1 effector. Similarly, the identification of mam was also likely facilitated by the fact that flies possess one mam gene, whereas mammals contain three mam orthologs. Thus, the study proposes that the comparative lack of genetic compensation/redundancy is an attractive advantage to Drosophila as a disease model system (Galindo, 2014).

The study suggests that further interrogation of MEF2 in RMS will open new avenues for RMS chemotherapy, which for high-risk disease has not improved for decades. For example, since MEF2 activity is tightly governed by class IIa histone deacetylases, histone deacetylase inhibitors are now ripe for preclinical testing as new anti-RMS agents. Additionally, it was found that the MEF2 cofactor Mastermind, which interacts with MEF2C and mediates crosstalk between Notch signals during myogenic differentiation, similarly influences PAX-FOXO1 pathogenicity in flies. Interestingly, Mastermind-specific, cell-permeable peptide inhibitors have been shown to block the progression of T-cell acute lymphoblastic leukemia in mice in vivo and thus are also new agents available for RMS preclinical testing. Further characterization of MEF2 in RMS cell and mouse models will continue to refine both our understanding and the potential targeting of MEF2 activity in RMS (Galindo, 2014).

In conclusion, the study postulates that: 1) The Drosophila PAX7-FOXO1 model is uniquely configured for the quick uncovering of new RMS genetic effectors with one simple genetic screening cross; 2) a putative PAX-FOXO1-to-MEF2/MASTERMIND axis underlies A-RMS; and 3) Drosophila conditional expression models are an efficient and powerful gene discovery platform for the rapid dissection of human disease (Galindo, 2014).


Agarwal, P., et al. (2011). The MADS box transcription factor MEF2C regulates melanocyte development and is a direct transcriptional target and partner of SOX10. Development 138(12): 2555-65. PubMed ID: 21610032

Anderson, C. M., Hu, J., Thomas, R., Gainous, T. B., Celona, B., Sinha, T., Dickel, D. E., Heidt, A. B., Xu, S. M., Bruneau, B. G., Pollard, K. S., Pennacchio, L. A. and Black, B. L. (2017). Cooperative activation of cardiac transcription through myocardin bridging of paired MEF2 sites. Development 144(7): 1235-1241. PubMed ID: 28351867

Apitz, H., Strunkelnberg, M., de Couet, H. G. and Fischbach, K. F. (2005). Single-minded, Dmef2, Pointed, and Su(H) act on identified regulatory sequences of the roughest gene in Drosophila melanogaster. Dev. Genes Evol. 215(9): 460-69. PubMed ID: 16096801

Apitz, H., Kambacheld, M., Höhne, M., Ramos, R. G. P., Straube, A. and Fischbach, K. F. (2004). Identification of regulatory modules mediating specific expression of the roughest gene in Drosophila melanogaster. Dev. Genes Evol. 214: 453-459. PubMed ID: 15278452

Artero, R., et al. (1998). The muscleblind gene participates in the organization of Z-Bands and epidermal attachments of Drosophila muscles and is regulated by Dmef2. Dev. Biol. 195(2): 131-143. PubMed ID: 9520330

Bagni, C., Bray, S., Gogos, J. A., Kafatos, F. C. and Hsu, T. (2002). The Drosophila zinc finger transcription factor CF2 is a myogenic marker downstream of MEF2 during muscle development. Mech. Dev. 117: 265-268. PubMed ID: 12204268

Bi, W., Drake, C. J. and Schwarz, J. J. (1999). The transcription factor MEF2C-null mouse exhibits complex vascular malformations and reduced cardiac expression of angiopoietin 1 and VEGF. Dev. Biol. 211(2): 255-67. PubMed ID: 10395786

Blais, A., et al. (2005). An initial blueprint for myogenic differentiation. Genes Dev. 19: 553-569. PubMed ID: 15706034

Bour, B. A., et al. (1995). Drosophila MEF2, a transcription factor that is essential for myogenesis. Genes Dev. 9: 730-741. PubMed ID: 7729689

Bryantsev, A. L., et al. (2012). Differential requirements for Myocyte Enhancer Factor-2 during adult myogenesis in Drosophila. Dev. Biol. 361(2): 191-207. PubMed ID: 22008792

Carvajal, J. J., Keith, A. and Rigby, P. W. (2008). Global transcriptional regulation of the locus encoding the skeletal muscle determination genes Mrf4 and Myf5. Genes Dev. 22(2): 265-76. PubMed ID: 18198342

Chang, C. I., et al. (2002). Crystal structures of MAP kinase p38 complexed to the docking sites on its nuclear substrate MEF2A and activator MKK3b. Molec. Cell 9: 1241-1249. PubMed ID: 12086621

Chen, S. L., et al. (2000). The steroid receptor coactivator, GRIP-1, is necessary for MEF-2C-dependent gene expression and skeletal muscle differentiation. Genes Dev. 14: 1209-1228. PubMed ID: 10817756

Chen, Z., Liang, S., Zhao, Y. and Han, Z. (2012). miR-92b regulates Mef2 levels through a negative-feedback circuit during Drosophila muscle development. Development 139: 3543-3552. Pubmed: 22899845

Clark, R. I., Tan, S. W., Pean, C. B., Roostalu, U., Vivancos, V., Bronda, K., Pilatova, M., Fu, J., Walker, D. W., Berdeaux, R., Geissmann, F. and Dionne, M. S. (2013). MEF2 is an in vivo immune-metabolic switch. Cell 155: 435-447. PubMed ID: 24075010

Cripps, M. R., et al. (1998). The myogenic regulatory gene Mef2 is a direct target for transcriptional activation by Twist during Drosophila myogenesis. Genes Dev. 12(3): 422-434. PubMed ID: 9450935

Cripps, R. M., Zhao, B. and Olson, E. N. (1999). Transcription of the myogenic regulatory gene Mef2 in cardiac, somatic, and visceral muscle cell lineages Is regulated by a Tinman-dependent core enhancer. Dev. Biol. 215(2): 420-430. PubMed ID: 10545248

Cunha, P. M., et al. (2010). Combinatorial binding leads to diverse regulatory responses: Lmd is a tissue-specific modulator of Mef2 activity. PLoS Genet. 6(7): e1001014. PubMed ID: 20617173

Damm C., et al. (1998). Independent regulatory elements in the upstream region of the Drosophila beta 3 tubulin gene (beta Tub60D) guide expression in the dorsal vessel and the somatic muscles. Dev. Biol. 199(1): 138-149. PubMed ID: 9676198

De Val, S., et al. (2004). Mef2c is activated directly by Ets transcription factors through an evolutionarily conserved endothelial cell-specific enhancer. Dev. Bio. 275: 424-434. PubMed ID: 15501228

de Velasco, B., Mandal, L., Mkrtchyan, M. and Hartenstein, V. (2006). Subdivision and developmental fate of the head mesoderm in Drosophila melanogaster. Dev. Genes Evol. 216(1): 39-51. PubMed ID: 16249873

Dichoso, D., et al. (2000). The MADS-box factor CeMEF2 is not essential for Caenorhabditis elegans myogenesis and development. Dev. Bio. 223: 431-440. PubMed ID: 10882527

Dodou, E., et al. (2004). Mef2c is a direct transcriptional target of ISL1 and GATA factors in the anterior heart field during mouse embryonic development. Development 131: 3931-3942. PubMed ID: 15253934

Duan, H., Skeath, J. B. and Nguyen, H. T. (2001). Drosophila Lame duck, a novel member of the Gli superfamily, acts as a key regulator of myogenesis by controlling fusion-competent myoblast development. Development 128: 4489-4500. PubMed ID: 11714674

Dunn, S. E., Chin, E. R. and Michel, R. N. (2000). Matching of calcineurin activity to upstream effectors is critical for skeletal mmuscle fiber growth. J. Cell Biol. 151: 663-672. PubMed ID: 11062266

Gajewski, K., et al. (1998). Combinatorial control of Drosophila mef2 gene expression in cardiac and somatic muscle cell lineages. Dev. Genes Evol. 208(7): 382-92. PubMed ID: 9732552

Gajewski, K., et al. (1999). The zinc finger proteins Pannier and GATA4 function as cardiogenic factors in Drosophila. Development 126: 5679-5688. PubMed ID: 10572044

Gajewski, K., et al. (2001). Pannier is a transcriptional target and partner of Tinman during Drosophila cardiogenesis. Dev. Bio. 233: 425-436. PubMed ID: 11336505

Galindo, K. A., Endicott, T. R., Avirneni-Vadlamudi, U. and Galindo, R. L. (2014). A rapid one-generation genetic screen in a Drosophila model to capture rhabdomyosarcoma effectors and therapeutic targets. G3 (Bethesda) 5(2):205-17. PubMed ID: 25491943

GarcĂ­a-Zaragoza, E., et al. (2008). CF2 activity and enhancer integration are required for proper muscle gene expression in Drosophila. Mech. Dev. 125: 617-630. PubMed ID: 18448314

Genikhovich, G. and Technau, U. (2011). Complex functions of Mef2 splice variants in the differentiation of endoderm and of a neuronal cell type in a sea anemone. Development 138(22): 4911-9. PubMed ID: 22007131

Gong, X., et la. (2003). Cdk5-mediated inhibition of the protective effects of transcription factor MEF2 in neurotoxicity-induced apoptosis. Neuron 38: 33-46. PubMed ID: 12691662

Gunthorpe, D., Beatty, K. E. and Taylor, M. V. (1999). Different levels, but not different isoforms, of the Drosophila transcription factor DMEF2 affect distinct aspects of muscle differentiation. Dev. Biol. 215(1): 130-45. PubMed ID: 10525355

Hadchoue, J., et al. (2003). Analysis of a key regulatory region upstream of the Myf5 gene reveals multiple phases of myogenesis, orchestrated at each site by a combination of elements dispersed throughout the locus. Development 130: 3415-3426. PubMed ID: 12810589

Han, A., et al. (2003). Sequence-specific recruitment of transcriptional co-repressor Cabin1 by myocyte enhancer factor-2. Nature 422(6933): 730-4. PubMed ID: 12700764

Han, T. H. and Prywes, R. (1995). Regulatory role of MEF2D in serum induction of the c-jun promoter. Mol. Cell. Biol. 15 (6): 2907-2915. PubMed ID: 7760790

Hayashi, S., Manabe, I., Suzuki, Y., Relaix, F. and Oishi, Y. (2016). Klf5 regulates muscle differentiation by directly targeting muscle-specific genes in cooperation with MyoD in mice. Elife [Epub ahead of print]. PubMed ID: 27743478

He, A., Kong, S. W., Ma, Q., and Pu, W. T. (2011). Co-occupancy by multiple cardiac transcription factors identifies transcriptional enhancers active in heart. Proc. Natl. Acad. Sci. 108: 5632-5637. PubMed ID: 21415370

He, J., Ye, J., Cai, Y., Riquelme, C., Liu, J. O., Liu, X., Han, A. and Chen, L. (2011). Structure of p300 bound to MEF2 on DNA reveals a mechanism of enhanceosome assembly. Nucleic Acids Res 39: 4464-4474. Pubmed: 21278418

Hinits, Y. and Hughes, S. M. (2007). Mef2s are required for thick filament formation in nascent muscle fibres. Development 134: 2511-2519. PubMed ID: 17537787

Huang, K., et al. (2000). Solution structure of the MEF2A-DNA complex: structural basis for the modulation of DNA bending and specificity by MADS-box transcription factors. EMBO J. 19: 2615-2628. PubMed ID: 10835359

Junion, G., et al. (2005). Mapping Dmef2-binding regulatory modules by using a ChIP-enriched in silico targets approach. Proc. Natl. Acad. Sci. 102(51): 18479-84. PubMed ID: 16339902

Kaushal, S., et al. (1994). Activation of the myogenic lineage by MEF2A, a factor that induces and cooperates with MyoD. Science 266: 1236-1240. PubMed ID: 7973707

Kato, Y., et al. (1997). BMK1/ERK5 regulates serum-induced early gene expression through transcription factor MEF2C. EMBO J. 16(23): 7054-7066. PubMed ID: 9384584

Kelly, K. K., Meadows, S. M. and Cripps, R. M. (2002). Drosophila MEF2 is a direct regulator of Actin57B transcription in cardiac, skeletal, and visceral muscle lineages. Mech. Dev. 110(1-2): 39-50. PubMed ID: 11744367

Krainc, D., et al. (1998). Synergistic activation of the N-methyl-D-aspartate receptor subunit 1 promoter by myocyte enhancer factor 2C and Sp1. J. Biol. Chem. 273(40): 26218-24. PubMed ID: 9748305

Kwon, C., Han, Z., Olson, E. N. and Srivastava. D. (2005). MicroRNA1 influences cardiac differentiation in Drosophila and regulates Notch signaling. Proc. Natl. Acad. Sci. 102(52): 18986-91. PubMed ID: 16357195

Laplante, M. and Sabatini, D. M. (2012). mTOR signaling in growth control and disease. Cell 149: 274-293. PubMed ID: 22500797

Lazaro, J.-B., Bailey, P. J. and Lassar, A. B. (2002). Cyclin D-cdk4 activity modulates the subnuclear localization and interaction of MEF2 with SRC-family coactivators during skeletal muscle differentiation. Genes Dev. 16: 1792-1805. PubMed ID: 12130539

Lee, Y., et al. (1997). Myocyte-specific enhancer factor 2 and thyroid hormone receptor associate and synergistically activate the alpha-cardiac myosin heavy-chain gene. Mol. Cell. Biol. 17: 2745-2755. PubMed ID: 9111345

Lilly, B., et al. (1994). D-MEF2: a MADS box transcription factor expressed in differentiating mesoderm and muscle cell lineages during Drosophila embryogenesis. Proc Natl Acad Sci 91: 5662-6. PubMed ID: 8202544

Lin, M.-H., et al. (1996). Myocyte-specific enhancer factor 2 acts cooperatively with a muscle activator region to regulate Drosophila tropomyosin gene muscle expression. Proc. Natl. Acad. Sci. 93: 4623-28. PubMed ID: 8643453

Lin, M.-H., et al. (1997). Ectopic expression of MEF2 in the epidermis induces epidermal expression of muscle genes and abnormal muscle development in Drosophila. Dev. Biol. 182: 240-255. PubMed ID: 9070325

Lin, S. C., et al. (1997a). PDP1, a novel Drosophila PAR domain bZIP transcription factor expressed in developing mesoderm, endoderm and ectoderm, is a transcriptional regulator of somatic muscle genes. Development 124(22): 4685-4696. PubMed ID: 9409684

Lin, S.- C. and Storti, R. V. (1997b). Developmental regulation of the Drosophila Tropomyosin I (TmI) gene is controlled by a muscle activator enhancer region that contains multiple cis-elements and binding sites for multiple proteins. Dev. Genet. 20(4): 297-306. PubMed ID: 9254904

Lin, Q., et al. (1998). Requirement of the MADS-box transcription factor MEF2C for vascular development. Development 125(22): 4565-74. PubMed ID: 9778514

Liotta, D., Han, J., Elgar, S., Garvey, C., Han, Z. and Taylor, M. V. (2007). The Him gene reveals a balance of inputs controlling muscle differentiation in Drosophila. Current Biol. 17: 1409-1413. PubMed ID: 17702578

Lovato, T. L., Benjamin, A. R. and Cripps, R. M. (2005). Transcription of Myocyte enhancer factor-2 in adult Drosophila myoblasts is induced by the steroid hormone ecdysone. Dev. Biol. 288(2): 612-21. PubMed ID: 16325168

Lu, J., et al. (2000). Regulation of skeletal myogenesis by association of the MEF2 transcription factor with class II histone deacetylases. Molec. Cell 6: 233-244

Liu, Z. P., et al. (2001). CHAMP, a novel cardiac-specific helicase regulated by MEF2C. Dev. Bio. 234: 497-509. PubMed ID: 11397016

Maeda, T., Chapman, D. L. and Stewart, A. F. (2002). Mammalian Vestigial-like 2, a cofactor of TEF-1 and MEF2 transcription factors that promotes skeletal muscle differentiation. J. Biol. Chem. 277(50): 48889-98. PubMed ID: 12376544

Mantrova, E. Y., Schulz, R. A. and Hsu, T. (1999). Oogenic function of the myogenic factor D-MEF2: Negative regulation of the Decapentaplegic receptor gene thick veins. Proc. Natl. Acad. Sci. 96: 11889-11894. PubMed ID: 10518546

Marco-Ferreres, R., et al. (2005). Co-operation between enhancers modulates quantitative expression from the Drosophila Paramyosin/miniparamyosin gene in different muscle types. Mech. Dev. 122(5): 681-94. PubMed ID: 15817225

Marin, M. C., Rodriguez, J. R. and Ferrus, A. (2004). Transcription of Drosophila troponin I gene is regulated by two conserved, functionally identical, synergistic elements. Mol. Biol. Cell 15: 1185-1196. PubMed ID: 14718563

Mas, J. A., Garcia-Zaragoza, E. and Cervera, M. (2004). Two functionally identical modular enhancers in Drosophila troponin T gene establish the correct protein levels in different muscle types. Mol. Biol. Cell 15: 1931-1945. PubMed ID: 14718560

McKinsey, T. A., Zhang, C. L., Lu, J. and Olson, E. N. (2000). Signal-dependent nuclear export of a histone deacetylase regulates muscle differentiation. Nature 408: 106-111. PubMed ID: 11081517

Michael, L. F., et al. (2001). Restoration of insulin-sensitive glucose transporter (GLUT4) gene expression in muscle cells by the transcriptional coactivator PGC-1. Proc. Natl. Acad. Sci. 98(7): 3820-5. PubMed ID: 11274399

Miska, E. A., et al. (1999). HDAC4 deacetylase associates with and represses the MEF2 transcription factor. EMBO J. 18: 5099-5107. PubMed ID: 10487761

Misquitta, L. and Paterson, B. M. (1999). Targeted disruption of gene function in Drosophila by RNA interference (RNA-i): A role for nautilus in embryonic somatic muscle formation. Proc. Natl. Acad. Sci. 96(4): 1451-6. PubMed ID: 9990044

Molkentin, J. D., et al. (1996a). MEF2B is a potent transactivator expressed in early myogenic lineages. Mol. Cell. Biol. 16: 3814-3824. PubMed ID: 8668199

Molkentin, J. D. and Olson, E. N. (1996b). Combinatorial control of muscle development by basic helix-loop-helix and MADS-box transcription factors. Proc. Natl. Acad. Sci. 93: 9366-73 . PubMed ID: 8790335

Morin, S., et al. (2000). GATA-dependent recruitment of MEF2 proteins to target promoters. EMBO J. 19: 2046-2055. PubMed ID: 10790371

Nakagawa, O., et al. (2005). Centronuclear myopathy in mice lacking a novel muscle-specific protein kinase transcriptionally regulated by MEF2. Genes Dev. 19: 2066-2077. PubMed ID: 16140986

Naya, F. J., et al. (1999). Transcriptional activity of MEF2 during mouse embryogenesis monitored with a MEF2-dependent transgene. Development 126(10): 2045-2052. PubMed ID: 10207130

Nguyen, H. T. and Xu, X. (1998). Drosophila mef2 expression during mesoderm development is controlled by a complex array of cis-acting regulatory modules. Dev. Biol. 204(2): 550-66. PubMed ID: 9882489

Nowak, S. J., Aihara, H., Gonzalez, K., Nibu, Y. and Baylies, M. K. (2012). Akirin links twist-regulated transcription with the Brahma chromatin remodeling complex during embryogenesis. PLoS Genet 8: e1002547. PubMed ID: 22396663

Okamoto, S.-I., et al. (2000). Antiapoptotic role of the p38 mitogen-activated protein kinase-myocyte enhancer factor 2 transcription factor pathway during neuronal differentiation. Proc. Natl. Acad. Sci. 97: 7561-7566. PubMed ID: 10852968

Pallavi, S. K., Ho, D. M., Hicks, C., Miele, L. and Artavanis-Tsakonas, S. (2012). Notch and Mef2 synergize to promote proliferation and metastasis through JNK signal activation in Drosophila. EMBO J. 31(13): 2895-907. PubMed Citation: 22580825

Park, M., Yaich, L. E. and Bodmer, R. (1998). Mesodermal cell fate decisions in Drosophila are under the control of the lineage genes numb, Notch, and sanpodo. Mech. Dev. 75(1-2): 117-26. PubMed ID: 9739121

Pena-Rangel, M. T., Rodriguez, I. and Riesgo-Escovar, J. R. (2002). A misexpression study examining dorsal thorax formation in Drosophila melanogaster. Genetics 160: 1035-1050. PubMed ID: 11901120

Penn, B. H., et al. (2004). A MyoD-generated feed-forward circuit temporally patterns gene expression during skeletal muscle differentiation. Genes Dev. 18: 2348-2353. PubMed ID: 15466486

Phan, D., et al. (2005). BOP, a regulator of right ventricular heart development, is a direct transcriptional target of MEF2C in the developing heart. Development 132(11): 2669-78. PubMed ID: 15890826

Postigo, A. A. and Dean, D. C. (1997). ZEB, a vertebrate homolog of Drosophila Zfh-1, is a negative regulator of muscle differentiation. EMBO J. 16(13): 3935-3943

Postigo, A. A., Ward, E., Skeath, J. B. and Dean, D. C. (1999). zfh-1, the Drosophila homologue of ZEB, is a transcriptional repressor that regulates somatic myogenesis. Mol. Cell. Biol. 19(10): 7255-63. PubMed ID: 10490660

Prokop, A., et al. (1996). Presynaptic development at the Drosophila neuromuscular junction: assembly and localization of presynaptic active zones. Neuron 17: 617-626

Pulipparacharuvil, S., et al. (2008). Cocaine regulates MEF2 to control synaptic and behavioral plasticity. Neuron 59(4): 621-33. PubMed ID: 18760698

Ramachandran, B., Yu, G. and Gulick, T. (2008). Nuclear respiratory factor 1 controls myocyte enhancer factor 2A transcription to provide a mechanism for coordinate expression of respiratory chain subunits. J. Biol. Chem. 283(18): 11935-46. PubMed ID: 18222924

Ranganayakulu, G., Zhao, B., Dokidis, A., Molkentin, J.D., Olson, E.N. and Schulz, R.A. (1995). A series of mutation in the D-MEF2 transcription factor reveal multiple functions in larval and adult myogenesis in Drosophila. Dev. Biol. 171: 169-181. PubMed ID: 7556894

Rushton, E., et al. (1995). Mutations in a novel gene, myoblast city, provide evidence in support of the founder cell hypothesis for Drosophila muscle development. Development 121: 1979-1988. PubMed ID: 7635046

Russo, S., et al. (1998). Myogenic conversion of NIH3T3 cells by exogenous MyoD family members: dissociation of terminal differentiation from myotube formation. J. Cell Sci. 111(6): 691-700. PubMed ID: 9471998

Sacilotto, N., et al. (2016) . MEF2 transcription factors are key regulators of sprouting angiogenesis. Genes Dev 30: 2297-2309. PubMed ID: 27898394

Sandmann, T., et al. (2006). A temporal map of transcription factor activity: Mef2 directly regulates target genes at all stages of muscle development. Dev. Cell 10(6): 797-807. PubMed ID: 16740481

Sandmann, T., et al. (2007). A core transcriptional network for early mesoderm development in Drosophila melanogaster. Genes Dev. 21: 436-449. PubMed ID: 17322403

Sartorelli, V., et al. (1997). Molecular mechanisms of myogenic coactivation by p300: Direct interaction with the activation domain of MyoD and with the MADS box of MEF2C. Mol. Cell. Biol. 17: 1010-26. PubMed ID: 9001254

Schlesinger, J., et al. (2011). The cardiac transcription network modulated by Gata4, Mef2a, Nkx2.5, Srf, histone modifications, and microRNAs. PLoS Genet. 7: e1001313. PubMed ID: 21379568

Schulz, R. A., et al. (1996). Expression of the D-MEF2 transcription in the Drosophila brain suggests a role in neuronal cell differentiation. Oncogene 12: 1827-1831. PubMed ID: 8622904

Simon, D. J., et al.. (2008). The microRNA miR-1 regulates a MEF-2-dependent retrograde signal at neuromuscular junctions. Cell 133(5): 903-15. PubMed ID: 18510933

Skerjanc, I. S., et al. (1998). Myocyte enhancer factor 2C and Nkx2-5 up-regulate each other's expression and initiate cardiomyogenesis in P19 cells. J. Biol. Chem. 273(52): 34904-10. PubMed ID: 9857019

Solera, C. and Taylor, M. V. (2009). The Him gene inhibits the development of Drosophila flight muscles during metamorphosis. Mech. Dev. 126: 595-603. PubMed ID: 19324085

Sparrow, B. D., et al. (1999). MEF-2 function is modified by a novel co-repressor, MITR. EMBO J. 18: 5085-5098. PubMed ID: 10487760

Spicer, D. B., et al. (1996). Inhibition of myogenic bHLH and MEF2 transcription factors by the bHLH protein Twist. Science 272: 1476-80. PubMed ID: 8633239

Spring, J., et al. (2002). Conservation of Brachyury, Mef2, and Snail in the myogenic lineage of jellyfish: A connection to the mesoderm of bilateria. Dev. Biol. 244: 372-384. PubMed ID: 11944944

Stronach, B. E., et al. (1999). Muscle LIM proteins are associated with muscle sarcomeres and require dMEF2 for their expression during Drosophila myogenesis. Mol. Biol. Cell 10: 2329-2342. PubMed ID: 10397768

Takeuchi, J. K., et al. (2005). Tbx20 dose-dependently regulates transcription factor networks required for mouse heart and motoneuron development. Development 132(10): 2463-74. PubMed ID: 15843409

Tanaka, K. K., Bryantsev, A. L. and Cripps, R. M. (2008). Myocyte enhancer factor 2 and chorion factor 2 collaborate in activation of the myogenic program in Drosophila. Mol. Cell Biol. 28: 1616-1629. PubMed ID: 18160709

Taylor, M. V., et al. (1995). Drosophila MEF2 is regulated by twist and is expressed in both the primordia and differentiated cells of the embryonic somatic, visceral and heart musculature. Mech. Dev. 50: 29-41. PubMed ID: 7605749

Taylor, M. V. (2000). A novel Drosophila, mef2-regulated muscle gene isolated in a subtractive hybridisation-based molecular screen using small amounts of zygotic mutant RNA. Dev. Biol. 220: 37-52. PubMed ID: 10720429

Thai, M. V., et al. (1998). Myocyte enhancer factor 2 (MEF2)-binding site is required for GLUT4 gene expression in transgenic mice. Regulation of mef2 DNA binding activity in insulin-deficient diabetes. J. Biol. Chem. 273(23): 14285-14292. PubMed ID: 9603935

Tokusumi, T., et al. (2007). U-shaped protein domains required for repression of cardiac gene expression in Drosophila. Differentiation 75: 166-174. PubMed ID: 17316386

von Both, I., et al. (2004). Foxh1 is essential for development of the anterior heart field. Dev. Cell 7: 331-345. PubMed ID: 15363409

Vrailas-Mortimer, A. D., Ryan, S. M., Avey, M. J., Mortimer, N. T., Dowse, H. and Sanyal, S. (2014). p38 MAP Kinase regulates circadian rhythms in Drosophila. J Biol Rhythms [Epub ahead of print]. PubMed ID: 25403440

Wang, D.-Z., et al. (2001). The Mef2c gene is a direct transcriptional target of myogenic bHLH and MEF2 proteins during skeletal muscle development. Development 128: 4623-4633. PubMed ID: 11714687

Wei, Q., Marchler, G., Edington, K., Karsch-Mizrachi, I. and Paterson, B. M. (2000). RNA interference demonstrates a role for nautilus in the myogenic conversion of Schneider cells by daughterless. Dev. Bio. 228: 239-255. PubMed ID: 11112327

West, A. G., Shore, P. and Sharrocks, A. D. (1997). DNA binding by MADS-box transcription factors: a molecular mechanism fo differential DNA bending. Mol. Cell. Biol. 17: 2876-87. PubMed ID: 9111360

White, K., et al. (1999). Microarray analysis of Drosophila development during metamorphosis. Science 286: 2179-2184. PubMed ID: 10591654

Wong, M. W., et al. (1994). Activation of Xenopus MyoD transcription by members of the MEF2 protein family. Dev Biol 166: 683-695. PubMed ID: 7813786

Wu, H., er al. (2000). MEF2 responds to multiple calcium-regulated signals in the control of skeletal muscle fiber type. EMBO J. 19: 1963-1973. PubMed ID: 10790363

Wu, H., et al. (2001). Activation of MEF2 by muscle activity is mediated through a calcineurin-dependent pathway. EMBO J. 20: 6414-6423. PubMed ID: 11707412

Yang, J., et al. (2002). Repression through a distal TCF-3 binding site restricts Xenopus myf-5 expression in gastrula mesoderm. Mech. Dev. 115: 79-89. PubMed ID: 12049769

Yang, S. H., Galanis, A. and Sharrocks, A. D. (1999). Targeting of p38 mitogen-activated protein kinases to MEF2 transcription factors. Mol. Cell. Biol. 19(6): 4028-38. PubMed ID: 10330143

Youn, H. D., Grozinger, C. M. and Liu, J. O. (2000a). Calcium regulates transcriptional repression of myocyte enhancer factor 2 by histone deacetylase 4. J. Biol. Chem. 275: 22563-22567. PubMed ID: 10825153

Youn, H. D. and Liu, J. O. (2000b). Cabin1 represses MEF2-dependent Nur77 expression and T cell apoptosis by controlling association of histone deacetylases and acetylases with MEF2. Immunity 13: 85-94. PubMed ID: 10933397

Youn, H.-D., Chatila, T. A. and Liu, J. (2000c). Integration of calcineurin and MEF2 signals by the coactivator p300 during T-cell apoptosis. EMBO J. 19: 4323-4331. PubMed ID: 10944115

Zetser, A., Gredinger, E. and Bengal E. (1999). p38 mitogen-activated protein kinase pathway promotes skeletal muscle differentiation. Participation of the Mef2c transcription factor. J. Biol. Chem. 274(8): 5193-200. PubMed ID: 9988769

Zhang, C. L., et al. (2001). Association of COOH-terminal-binding protein (CtBP) and MEF2-interacting transcription repressor (MITR) contributes to transcriptional repression of the MEF2 transcription factor. J. Biol. Chem. 276(1): 35-9. PubMed ID: 11022042

Zhang, C. L., et al. (2002). Class II histone deacetylases act as signal-responsive repressors of cardiac hypertrophy. Cell 110: 479-488. PubMed ID: 12202037

Zhao, M., et al. (1999). Regulation of the MEF2 family of transcription factors by p38. Mol. Cell. Biol. 19(1): 21-30. PubMed ID: 9858528

Zhao, X., et al. (2005). Regulation of MEF2 by histone deacetylase 4- and SIRT1 deacetylase-mediated lysine modifications. Mol. Cell. Biol. 25(19): 8456-64. PubMed ID: 16166628

Myocyte enhancer factor 2: Biological Overview | Evolutionary Homologs | Regulation | Targets of Activity and Protein Interactions | Developmental Biology | Effects of Mutation

date revised: 22 May 2017

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