Although Merlin and Moesin proteins are frequently coexpressed in developing tissues, they display distinct subcellular localizations. In polarized epithelia, such as the embryonic hindgut, salivary gland, or the imaginal disc, both Moesin and Merlin are found in the highest concentration in the most apical part of the cell. Colocalization experiments in wing imaginal discs demonstrate that at least part of the detected Merlin and Moesin protein is associated with the adherens junction as is the beta-Catenin homolog, Armadillo. Moesin is localized to the apical cap of imaginal disc cells, a region known to contain abundant microvilli. Both Moesin and filamentous actin are localized in microvilli present during cellularization in the preblastoderm embryo. Merlin does not localize with actin or Moesin in apical buds. While Moesin is observed in continuous association with the plasma membrane, as is typical for an ERM family protein, Merlin is found in punctuate structures at the membrane and in the cytoplasm. Investigation of Merlin distribution in cultured cells demonstrates that it is associated with endocytic compartments. Merlin may not interact with cytoskeletal actin in the same way that has been proposed for other members of the ERM family. It is thought that Merlin is associated with membrane internalized from the cell surface. As a result of these studies, it is proposed that whereas Moesin may fulfill all the functions of vertebrate ERM proteins, Merlin protein has unique functions in the cell which differ from those of other ERM family members, perhaps being involved in endocytosis. It is concluded that Merlin is a functionally distinct ERM family member (McCartney, 1996).

In contrast to the largely complementary pattern observed for Merlin and Moesin during oogenesis, the two appear to be coexpressed in most cells during embryogenesis. Merlin and Moesin are present from cellularization throughout embryonic development. Merlin expression is found to be enhanced in the early mesoderm of the germband extended embryo, whereas Moesin is expressed uniformly throughout the embryo at this stage. Late in embryogenesis, both proteins are expressed ubiquitously throughout the tissues of the embryo, including the epidermis, salivary glands, foregut, midgut, hindgut, and the embryonic nervous system. At this stage Merlin expression is enhanced in the midgut (McCartney, 1996).

In the embryonic central nervous system (stage 15), Merlin and Moesin staining is detected in the neuropil, a structure composed of the developing axonal bundles of the ventral nerve cord and in the developing brain. Merlin and Moesin localization is also observed in the neuronal cell bodies of the CNS, with Moesin expression enhanced in these tissues. During stage 11 of embryogenesis, the neurons of the embryonic peripheral nervous system develop from specialized regions within the epidermis termed the proneural clusters. The cell bodies of the differentially bipolar sensory neurons can be observed in a regular pattern within the epidermis of a stage 17 embryo. At this stage, Moesin is enriched at the membranes of these cells, whereas Merlin is found to localize to an intensely staining spot within the cell body (McCartney, 1996)

Mammalian Ezrin, Radixin and Moesin (ERM) are components of the cortical actin cytoskeleton and membrane processes such as filopodia and microvilli. Their C-terminal tails contain an extended region that is predicted to be helical, an actin binding domain, and a region(s) that participates in self-association. An in vivo fluorescent actin binding protein (GFP-Moe) as engineered by joining sequences that encode the jellyfish green fluorescent protein (GFP) to sequences that encode the C-terminal end of the sole Drosophila ERM homolog, Moesin (Moesin gene product). Transgenic flies expressing this fusion protein under control of the hsp70 promoter were generated and used for analysis of cell shape changes during morphogenesis of various developmental stages and tissues. Following heat shock, high levels of stable fusion protein are produced by all somatic tissues. GFP-Moe localizes to the cortical actin cytoskeleton, providing a strong in vivo marker for cell shape and pattern during epithelial morphogenesis. The protein also becomes highly enriched in pseudopods, microvilli, axons, denticles, the border cell process, and other membrane projections, potentially by binding to endogenous Moesin as well as actin. GFP-Moe can be used to examine the development and behavior of these dynamic structures in live specimens. A bright green fluorescent, presumably actin-rich, polar cell proboscis is observed that inserts itself into the forming micropyle and appears to maintain an opening for sperm passage around which the chorion is formed. The existence of an actin-rich purse string is confirmed at the leading edge of the lateral epidermis and a dynamic analysis is provided of its behavior as it migrates during dorsal closure. Observations of embryos, larvae, and pupae show that GFP-Moe is also useful for labeling the developing nervous system and will be a good general marker of dynamic cell behavior during morphogenesis in live tissues and demonstrate that fusion of a subcellular localization signal to GFP greatly increases its utility as a cell marker (Edwards, 1997).

Larval and Pupal

In the developing eye Moesin is localized at the membranes of the cone cells, secondary and tertiary pigment cells, and in the bristle precursor cells. In contrast, Merlin is localized primarily in the cytoplasm of the secondary and tertiary pigment cells, and is greatly enhanced in the bristle precursor cells, which are interspersed between outer pigment cells. Both Merlin and Moesin are more intensely expressed in the center of each ommatidium. This corresponds to the region of the rhabdomeres, the photosensitive microvilli of the photoreceptors, which project into the center of each ommatidium (McCartney, 1996).

Fine details of the developing pupal CNS and visual system are vividly highlighted by the GFP-Moe. In addition to the rhabdomeres and/or cell bodies of the photoreceptors, axons are labeled as they project in an ordered fashion into the lamina and then to the optic lobes. Thoracic and abdominal neuromeres are apparent. In the retina, the arrangement of ommatidia and their constituent cells can be seen. The presumptive margin of the wing is revealed by a double row of bristle precursors. In later stage female pupae, the somatic cells of the ovary, especially the terminal filaments, are prominent. Some muscles show a striated GFP signal, though it is weaker and more variable than one might expect given the quantity of actin present. The indirect flight muscles that pack the thorax accumulate very little fusion protein and no striations are evident. It is suspected that the moesin binding site on actin is occluded to a greater or lesser extent in different muscle types (Edwards, 1997).


During oogenesis, Merlin and Moesin display strikingly different tissue distributions: this distinction is clearly observed as early as the germarium, the location of the germline stem cells. Merlin is expressed predominantly in the germline, while Moesin is expressed at greater levels in the follicle cells. In addition, Merlin expression becomes enhanced in the developing oocyte at approximately stage 6 of development and persists until the end of oogenesis. Lower levels of Merlin expression are also detected at the apical ends of the follicle cells at stage 10, late in oogenesis. In contrast, Moesin expression is found at the apical and basolateral ends of the follicular epithelium, although some expression is detected in the germline of early egg chambers and in the nurse cells at stage 10. Fully developed oocytes (stage 14) clearly display membrane associated Merlin, while no Moesin expression is detected at this stage (McCartney, 1996).

The GFP-Moe fusion protein uniformly outlines the somatic follicle cells as they encase individual germ cell cysts in the germarium and subsequently form egg chambers separated by interfollicular stalks. GFP signal is most intense at the apical (inner) ends of the follicle cells, but the signal is not polarized in the interfollicular stalk cells. Germ cells are relatively devoid of signal, apparently because the heat shock promoter is, in this case, not strongly inducible in the germline. With high gain, a faint signal is detectable at the nurse cell membranes and the ring canals, the actin-rich structures lining the passages between the germ cells. In stage 10 egg chambers, most follicle cells belong to two populations: the oocyte follicle cells, which form a columnar epithelium over the oocyte, and the nurse cell follicle cells, which flatten and spread over the adjoining nurse cells. From the basal (outer) ends of the oocyte follicle cells protrude extremely long microvilli that exhibit bright fluorescence. Comparable projections cannot be seen by phalloidin staining in wild-type chambers, suggesting that they may actually be induced by expression of the fusion protein. The apical ends also show microvilli, but they are much shorter than the basal projections and indistinguishable from microvilli seen in wild-type phalloidin- stained chambers. GFP signal is not seen at the lateral membranes. In contrast to the oocyte follicle cells, projections are not visible on the nurse cell follicle cells, and on earlier stage follicle cells only short, sparse projections appear (Edwards, 1997).

A pair of cells at each end of the newly formed egg chamber, called polar follicle cells (PFCs), are specified to differentiate from the rest of the follicular epithelium. The PFCs express characteristic marker proteins, adopt specific morphologies, and cease proliferation. This last property allows the specific labelling of these cells by transiently inducing hs-GFP-Moe. One day after induction, the GFP signal in the PFCs remains strong; the signal in the other follicle cells is diluted by proliferation. Two days after induction, the PFCs are labeled with high specificity. Stalk cells also stop dividing and remain labeled (Edwards, 1997).

While most follicle cells move posteriorly over the growing oocyte, approximately eight cells (border cells) at the anterior tip of the egg chamber delaminate from the follicular epithelium. They migrate between the nurse cells toward the anterior face of the oocyte, remaining tightly adherent to each other as they crawl. After arriving at the oocyte, they are met by centripetally migrating follicle cells and collaborate with them to form the micropyle, the sperm entry port in the mature eggshell. The anterior PFCs, observed using GFP-Moe, are invariably included in the border cell cluster. In fact, the PFCs are always found in the same position, central in the cluster. This suggests that they have a key role in organizing the border cells. The PFCs play another important role after the border cells arrive at the oocyte. The PFC touches the oocyte with a broad cell surface projection. As the micropyle is built, the projection extends and narrows to form the border cell process. This process penetrates the micropyle, creating the central canal through which the sperm enters. At all stages this projection is labeled even intensely than the cell body. These observations suggest that the anterior PFCs are fated early in oogenesis to eventually form the micropylar canal. Since there are many cells in the region secreting autofluorescent chorion material, this provides a unique and powerful method for following this important developmental process in living specimens. The results with live specimens agree strongly with accounts of micropyle formation suggesting that the fusion protein does not interfere with cell behavior. Interestingly, the border cell process is rich in microtubules, suggesting that its morphogenesis may be dependent on both actin- and microtubule-dependent processes (Edwards, 1997).

Effects of Mutation or Deletion

Moesin is required for Oskar anchoring

In Drosophila, development of the embryonic germ cells depends on posterior transport and site-specific translation of oskar (osk) mRNA and on interdependent anchoring of the osk mRNA and protein within the posterior subcortical region of the oocyte. Transport of the osk mRNA is mediated by microtubules, while anchoring of the osk gene products at the posterior pole of the oocyte is suggested to be microfilament dependent. To date, only a single actin binding protein (TropomyosinII) has been identified with a putative role in osk mRNA and protein anchoring. Mutations in the Drosophila moesin-like (moe) gene, which encodes another actin binding protein, result in delocalization of osk mRNA and protein from the posterior subcortical region and, as a consequence, in the failure of embryonic germ cell development. In moe mutant oocytes, the subcortical actin network is detached from the cell membrane, while the polarized microtubule cytoskeleton is unaffected. Colocalization of ectopic actin and Osk protein in moe mutants suggests that the actin cytoskeleton anchors Osk protein to the subcortical cytoplasmic area of the Drosophila oocyte (Jankovics, 2002).

Moe is the Drosophila member of the ERM protein family, which contains three vertebrate members: ezrin, radixin, and moesin. ERM proteins have a C-terminal actin binding domain and an N-terminal FERM domain, which is responsible for interaction with several membrane proteins. Based on their structure and predominant subcortical distribution, ERM proteins have been suggested to function as crosslinkers between the cell membrane and the actin cytoskeleton. ERM proteins have been demonstrated to take part in several biological processes, such as cell-cell adhesion, maintenance of cell shape, cell motility, and internal membrane trafficking. Despite their distinct expression pattern, the vertebrate ERM protein family members show functional redundancy. The Drosophila genome, however, contains only a single ERM homolog: Moesin. Drosophila is a valuable model organism for studying ERM functions, since the ERM homolog is not genetically redundant (Jankovics, 2002).

In a screen for P element-induced maternal effect germ cell-less mutations, a recessive allele of the moe gene was identified. The moeGT193 allele was isolated by making use of a newly designed EGFP-GT mutator P element. EGFP-GT was made by replacing the Gal4 marker gene of a dual-tagging gene trap element, pGT1, with the EGFP marker gene. A collection of 200 viable, X-chromosomal EGFP-GT insertions was generated by the attached-X technique and was screened for maternal effect germ cell-less phenotype by hand dissection of adult test animals. The moeGT193 allele was identified as a weak maternal effect germ cell-less allele. Subsequent complementation analyses with P element-induced mutations obtained from the Bloomington Stock Center revealed the existence of five additional insertional alleles of moe. Combinations of different moe alleles resulted in germ cell-less phenotypes with a penetrance of 1%-61%. For further analyses, the moeEP1652/moeG0415 and moeEP1652/Df(1)KA14 combinations were chosen; these resulted in 60% and 61% germ cell-less phenotypes, respectively. Hereafter, the phenotype of these combinations will be referred to as the moe mutant phenotype (Jankovics, 2002).

Complementation analyses revealed several additional pleiotropic moe phenotypes such as lethality, female sterility, and imperfect eye and wing development. Western blot analysis of moe mutant ovaries revealed a reduction in Moe protein levels, while precise excisions of P elements from moe alleles restored wild-type protein levels and germ cell development. These results demonstrate that the maternal effect germ cell-less phenotype is a direct consequence of P element insertions in the moe gene (Jankovics, 2002).

Since ERM proteins have been suggested as functioning as crosslinkers between the actin network and the cell membrane, the organization of actin filaments was examined in moe oocytes. Instead of a tight, wild-type subcortical localization, in moe mutants the actin network seems to be detached from the cortex. It intrudes into the cytoplasm of the oocyte either at the posterior pole or at more lateral regions. Interestingly, mislocalized subcortical actin is also found in Schizosaccharomyces pombe after being transformed with truncated Drosophila moe cDNA. In this heterologous transformation experiment, actin abnormalities were also coupled with cell shape changes. This raises the possibility that observed actin abnormalities in Drosophila oocytes may be the result of abnormally shaped oocytes. To rule out this possibility, the cell and vitelline membranes were visualized by making use of fluorescently labeled lectins (Lycopersicon esculentum and Datura stramonium lectins, respectively) and immunostained for DE-cadherin, a transmembrane protein that is present in all cell membranes of egg primordia. In developing mutant eggs, however, normal distribution of the lectins and DE-cadherin stainings are detected. Furthermore, by simultaneous visualization of the cell membrane-specific lectin and actin, it has been shown that, at actin intrusion sites, the cell membrane is normal. These results demonstrate that, in moe mutants, the shape of the oocyte is normal and the observed actin abnormality is due to detachment of subcortical actin from the cell membrane. It is concluded, therefore, that one of the functions of moe in developing oocytes is to crosslink the subcortical actin network and the cell membrane (Jankovics, 2002).

Embryonic germ cell formation is initiated during oogenesis by posterior localization of a highly specialized cytoplasmic region, the pole plasm. Since the assembly of the pole plasm depends on the anterior-to-posterior transport and subsequent anchoring of osk mRNA, the distribution of osk mRNA was examined in moe oocytes. Instead of the characteristic wild-type posterior localization, several different types of abnormal osk mRNA distribution were found in the mutants. Most frequently, the mislocalized osk mRNA appears in a scattered pattern concentrated near the posterior pole. Mislocalization of osk mRNA to the central regions of oocytes was also observed. In some stage-10 oocytes, ectopic osk mRNA was found to be localized tightly to the lateral cell cortex. In order to gain insight into the time course of osk mRNA localization in mutants, the penetrance of these phenotypes was measured and compared in stages 9 and 10. A slight, but convincing, decrease of the wild-type osk mRNA localization was observed in stage 10, and this decrease suggests that moe mutations do not interfere with the early posterior transport of osk mRNA but rather with its anchoring to the posterior pole. Another pole plasm component, the Staufen (Stau) protein, which always colocalizes with osk mRNA, consistently shows a distribution identical to that of osk mRNA in mutant oocytes (Jankovics, 2002).

Since correct localization of osk mRNA requires proper functioning of both the oocyte and follicle cells, and moe is expressed predominantly in the follicle cells, moe mutant germline clones were generated to identify the cell type in which moe mutations exert their effect. In developing eggs composed of the homozygous germline and heterozygous follicle cells for moeG0415, moeG0404, moeG0067, and moeG0323 mutations, osk mRNA mislocalization phenotypes were found to be identical to those of moeEP1652/DmoeG0415and moeEP1652/Df(1)KA14 mutant oocytes. This result demonstrates that moe is required in the germ cells for posterior localization of osk mRNA (Jankovics, 2002).

To investigate whether moe mutations affect other localized determinants in the Drosophila oocyte, the distributions of gurken (grk) and bicoid (bcd) mRNAs were examined. In mutant oocytes, normal grk and bcd mRNA localization was found, which demonstrates that the moe mutant phenotype is osk specific. Similar to moe mutations, par-1 and Rab11 mutant alleles cause abnormal osk mRNA localization and normal distribution of the bcd and grk mRNAs. In these mutants, a plus end microtubule marker molecule, Kin:β-Gal, is localized to the central region of the oocyte, while the minus end-specific Nod:β-Gal remains at the anterior pole. In contrast to these mutants, however, normal Nod:β-Gal and Kin:β-Gal localization was observed in moe oocytes. Furthermore, a normal arrangement of the microtubule network was revealed both by immunostaining and by direct in vivo visualization of the microtubules by using Tubulin:GFP or Tau:GFP fusion proteins. By a simultaneous visualization of STAU protein and microtubule plus ends, Stau mislocalization was shown not to be a consequence of an abnormal microtubule network. In moe mutant oocytes, the plus ends of the microtubules normally point to the posterior pole, as demonstrated by the correct localization of Kin:β-Gal. In contrast, however, Stau is mislocalized, suggesting again that the mislocalization of osk mRNA is not a consequence of abnormal transport; rather, it appears to be caused by abnormal anchoring (Jankovics, 2002).

In order to further investigate the role of Moe in osk regulation, the level and distribution of Osk protein was examined in moe oocytes. In mutant ovaries, a reduced level of Osk was found by Western analysis. Consistent with this observation, no Osk was detected by immunostaining in the majority of the mutant oocytes, while, in the remaining cases, the pattern of the mislocalized Osk protein was found in a spatial and temporal distribution similar to that of osk mRNA. Osk was mostly found in a scattered pattern at the posterior region of the oocytes. In the majority of moe oocytes in which Osk was delocalized, the actin network appears to be attached normally to the cell membrane, at least as far as can be visualized with light microscopy. This indicates that moe might have a role in Osk anchoring, which is separable from its actin-cell membrane crosslinking function (Jankovics, 2002).

In the rare cases when detachment of actin from the cell membrane occurs at the posterior pole, it was observed that Osk protein colocalizes with the ectopic actin network. This ectopic colocalization of the subcortical actin and Osk protein in moe mutants reveals that the actin network has Osk anchoring capacity and strongly suggests that the actin network anchors Osk at the normal place, at the posterior subcortical region, too. In moe mutants, the abnormal osk mRNA and protein localization phenotype is observed when osk products are tightly localized to lateral segments of the subcortical actin. These results confirm a model that proposes that the focused posterior localization of osk is not defined by a special segment of the posterior subcortical actin; rather, it is determined by the microtubule-based posterior transport of osk mRNA to this region (Jankovics, 2002).

Thus, partial loss of moe activity weakens the ability of the actin network to anchor osk mRNA and protein, resulting in different degrees of their delocalization. moe has actin-cell membrane crosslinking activity in the developing oocyte, and evidence is presented that factors other than actin define the site of osk mRNA and protein localization (Jankovics, 2002).

Dmoesin controls actin-based cell shape and polarity during Drosophila melanogaster oogenesis

Ezrin, Radixin and Moesin (ERM) proteins are thought to constitute a bridge between the actin cytoskeleton and the plasma membrane (PM). This study reports a genetic analysis of Drosophila Moesin, the sole member of the ERM family in Drosophila. Moesin is required during oogenesis for anchoring microfilaments to the oocyte cortex. Alteration of the actin cytoskeleton resulting from Moesin mutations impairs the localization of maternal determinants, thus disrupting antero-posterior polarity. This study also demonstrates the requirement of Moesin for the specific organization of cortical microfilaments in nurse cells and, consequently, mutations in Moesin produce severe defects in cell shape (Polesello, 2002).

Actin-based cellular processes depend on actin filament polymerization and branching and must involve a tight control of microfilament attachment to the PM. Ezrin, Radixin and Moesin (ERM) are highly homologous proteins that were originally discovered in mammals and which accumulate in numerous actin-rich structures. ERM proteins have been shown to bind to both F-actin and membrane proteins, and thus could constitute a bridge between the actin cytoskeleton and the cell membrane (Polesello, 2002).

In ERM proteins, a central alpha-helical region separates two domains: the amino-terminal FERM domain, a membrane-linking module found in several other proteins (including the tumour suppressor Merlin/neurofibromatosis 2 (NF2) and the F-actin binding site composed of the 34 carboxy-terminal amino acids). ERM proteins exist in a dormant state, in which the FERM domain interacts with the C-terminal tail, resulting in a closed configuration. Activation of ERM results in the opening of the molecular structure, thus unmasking the two protein domains, which can then interact with their respective partners. Activation of ERM also correlates with the phosphorylation of a critical threonine residue located in the F-actin binding site and several signalling pathways are candidates for this phosphorylation event (Polesello, 2002 and references therein).

ERMs are involved in the control of actin-based morphogenesis, as their inactivation -- either by antisense oligonucleotides or laser irradiation -- alters the shape and adhesive properties of cultured cells. However, targeted disruption of the mouse Moesin gene does not result in any abnormal phenotype. Therefore, although accumulated data suggest that ERMs are important regulators of actin-based cellular processes, a genetic system to allow analysis of ERM functions during development has been lacking (Polesello, 2002 and references therein).

Mutant alleles in the single Drosophila ERM gene, Moesin have been isolated, its function during oogenesis has been analyzed. Evidence is presented that Moesin is required for the antero-posterior polarity of the oocyte. Mutations in Moesin disrupt the anchorage of F-actin to the oocyte cortex and impair the localization of posterior determinants, resulting in the absence of embryonic posterior structures. Moesin accumulates specifically at the cortex of nurse cells, where it colocalizes with a dense mesh of microfilaments. Alteration of Moesin generates marked defects in this polarized F-actin network, resulting in abnormal cell shape. To study the role of the conserved C-terminal Thr 559 residue in vivo, the consequences of mutations that prevent, or mimic, its phosphorylation has been analyzed. These alterations modify the subcellular localization of Moesin. Furthermore, these substitutions cause defects in the organization of the polarized actin cytoskeleton and also affect the distribution of posterior determinants (Polesello, 2002).

Three P-element insertions within the Dmoesin locus have been isolated and characterized. All the mutations belong to a single complementation group and the lethality of each mutant is a result of the P-element insertion. In the DmoePG26 mutant, the insertion allows the expression of Gal4 under the control of Moe promoter elements. When Moe cDNA was introduced into DmoePG26 flies downstream of Gal4 binding sites (upstream activating sequence), viability was fully restored and all mutant phenotypes were suppressed. The accumulation of Moe mRNA is strongly reduced in mutants, confirming that they are, if not nulls, then strong Moesin alleles. Genetic tests show that Dmoe106 and Dmoe54 are stronger alleles than DmoeX5 and DmoePG26, respectively. These results indicate that the isolated mutations specifically alter Moesin and show that Moesin is an essential Drosophila gene (Polesello, 2002).

A first indication of the developmental functions of Moesin was provided by the observation that a proportion (7%-11%) of the eggs laid by heterozygous Moe females display segmentation defects. These abnormal embryos (referred to hereafter as Moe embryos) lack posterior structures. This so-called 'posterior group' phenotype is similar to the phenotype of mutations in maternal genes that determine antero-posterior polarity, suggesting that the early steps of segmentation are altered in Moe embryos. Accordingly, Wingless (Wg), which is normally expressed in 14 segmental stripes, is instead localized to a few wide bands in the trunk of Moe embryos, attesting to the absence and fusion of segments. These segmentation anomalies are observed independently of the zygotic genotype of the embryos, confirming that they result from the reduction of maternal Moe. Furthermore, Moe embryos produce only a few posterior pole cells, which are the germ cell precursors. The posterior region of early Drosophila embryos contains the pole plasm, which includes Vasa and Nanos proteins responsible for the formation of pole cells and posterior segments, respectively. The posterior accumulation of both Vasa protein and nanos mRNA is strongly reduced in Moe embryos, whereas Bicoid protein is localized normally in an anterior gradient. Therefore, these results show that a reduction of maternal Moe specifically affects the early organization of the posterior pole (Polesello, 2002).

The Drosophila antero-posterior axis is established during oogenesis, where the localized activity of Oskar determines the posterior pole and recruits Nanos and Vasa. To assay the role of Moe in posterior pole formation, Moe germline mutant clones were generated. Females with a Moe mutant germline are sterile and do not lay eggs. In wild-type oocytes, Oskar accumulates in a crescent that is tightly localized to the posterior of the oocyte cortex. By contrast, in Moe oocytes, Oskar is found at more anterior locations and forms abnormal fibers, apparently loosely bound to the cortex. Localization of the Oskar protein is linked to the polarized accumulation of oskar mRNA. oskar is normally transcribed in Moe oocytes and until stage-8, its localization is indistinguishable from that in wild-type oocytes. However, the posterior localization of oskar mRNAs that starts from stage-9 is affected in Moe mutants. Whereas oskar mRNAs are restricted to the posterior of wild-type oocytes, a diffuse signal is seen throughout the oocyte cytoplasm (ooplasm) in stage-10 Moe oocytes. In some cases, a weak accumulation of oskar mRNAs is observed at the posterior pole, indicating that Moe mutations do not completely abrogate the transport of oskar mRNAs. In contrast, gurken and bicoid mRNAs localize normally at an antero-dorsal position and the anterior margin, respectively. This indicates that Moe mutations do not alter the formation of the dorso-ventral axis or the anterior pole. Localization of Oskar depends on Staufen, an RNA-binding protein that is required for the transport of oskar mRNA. Compared with wild-type oocytes, the posterior accumulation of Staufen is reduced in Moe oocytes; Staufen is not firmly localized to the posterior and a diffuse staining is observed in the ooplasm (Polesello, 2002).

The establishment of oocyte polarity and Oskar localization depends on microtubule organization. To determine whether Moe mutations affect microtubules, the movements of yolk granules were analyzed in living egg chambers. In wild-type oocytes, granules move erratically up to stage-10a and eventually flow rapidly in the coordinated cytoplasmic streaming. These movements are blocked by microtubule-depolymerizing drugs, and mutations affecting microtubule organization cause premature streaming. No sign of premature cytoplasmic streaming was observed in Dmoe106 germline clones, although typical coordinated movements of granules are observed at stage-10b-11. This indicates that Moe mutations do not disrupt the microtubule cytoskeleton. Microtubule polarity was analyzed using Nod-ßgalactosidase, which localizes to the minus-end of microtubules. As in the wild-type oocytes, Nod-ßgalactosidase accumulates in the anterior of the mature Moe oocyte. Finally, a lacZ fusion to Kinesin, a plus-end-directed motor that is required for the localization of posterior determinants, was examined. From late-stage-8 onwards, Kinesin-ßgalactosidase displays a polarized distribution in wild-type oocytes. It accumulates transiently (stage 9-10) at the posterior extremity of the oocyte, where it colocalizes with Staufen. In DmoeX5 germline clones, Kinesin-ßgalactosidase localizes normally in stage-9 oocytes. At stage 10a, the staining, although reduced, was still detected at the posterior end, confirming the normal polarity of microtubules in DmoeX5 oocytes. However, Staufen does not completely colocalize with Kinesin-ßgalactosidase in DmoeX5 oocytes; Staufen forms ectopic clumps and displays diffuse staining in the ooplasm. This further supports the conclusion that the mislocalization of posterior determinants observed in Moe oocytes does not result from the disruption of early microtubule nucleation and polarity. Taken together, these results indicate that Moesin is specifically required for the localization of posterior determinants and suggest a critical role of Moe in the functional organization of the oocyte cortex (Polesello, 2002).

To analyse the role of Moe in the organization of the oocyte cortex, the localization of a green fluorescent protein (GFP)-tagged Moesin protein, which fully rescues DmoePG26 mutant phenotypes, was examined. From the early stages of oogenesis until the development of mature egg chambers, Moesin-GFP is present in the oocyte cytosol and accumulates at the cortex, where it colocalizes with F-actin. This membrane-associated fraction corresponds to activated Moesin, because it is recognised by an antibody specific for activated ERM. The consequences of Moe mutations on the cortical actin cytoskeleton were analzed. In the wild-type oocyte, microfilaments accumulate in the cortex and form bundles parallel to the membrane. In oocytes from DmoeX5 germline clones, microfilaments appear loosely bound to the posterior and lateral cortex. In stronger Dmoe alleles, in addition to filaments loosely attached to the membrane, packs of F-actin are also found in the ooplasm. This raises the possibility that these actin defects are responsible for the altered posterior determinant distribution. As observed at earlier stages, cortical F-actin has an abnormal fibrous appearance in stage-10 Dmoe oocytes, and clumps of F-actin are also observed in the ooplasm. Interestingly, Oskar associates with these abnormal actin structures, colocalizing with both lateral F-actin fibres and ooplasmic clumps. These data show that Oskar closely interacts with F-actin and strongly support the conclusion that the mislocalization of Oskar observed in Dmoe oocytes results from defects of the actin cytoskeleton (Polesello, 2002).

Nevertheless, oocyte polarity depends on a signal from the posterior follicle cells, and Dmerlin, which is structurally similar to Moesin, is required in the posterior follicle cells to ensure oocyte polarity. Thus, whether Dmoe activity is also required in posterior follicle cells to organize oocyte polarity was analyzed. When only follicle cells are mutant for Dmoe, neither alteration of the oocyte F-actin organization, nor mislocalization of Oskar was observed. Altogether, these results show that Dmoe is required cell-autonomously in the germline, to anchor actin microfilaments to the oocyte cortex and to localize posterior determinants (Polesello, 2002).

Phosphorylation of a threonine residue located in the actin-binding domain has been shown to regulate ERM activity in mammalian cells. To assay the role of this conserved residue for Drosophila Moesin function, transgenes carrying a Thr 559 mutation that either prevents (Thr-Ala; TA) or mimics (Thr-Asp; TD) phosphorylation of ERM33 were generated. These Moesin variants were fused to GFP and expressed specifically in the germline. Unlike wild-type Moesin-GFP, the entire pool of phosphomimetically mutated protein is directed to the membrane, where it colocalizes with F-actin. Although slightly enriched in the cytoplasm, the non-phosphorylatable form of Moesin associates with the oocyte membrane, where it accumulates ectopically at the anterior. Expression of Moesin-TA also results in the formation of F-actin clumps in the ooplasm. These results show that Thr 559 is involved in the regulation of the subcellular distribution of Moesin and identify its importance for the organization of the oocyte actin cytoskeleton. Consistently, neither Moesin-TD nor Moesin-TA can substitute for wild-type Moesin in rescue assays. In addition, whereas overexpression of MoesinWT-GFP does not alter the distribution of Oskar, the posterior localization of Oskar is either undetectable or strongly reduced in oocytes that express Moesin-TA. By contrast, expression of Moesin-TD results in a marked over-accumulation of Oskar at the posterior end of the oocyte. These data show that Moesin must be properly regulated to allow the normal localization of Oskar at the posterior end of the oocyte (Polesello, 2002).

Although the oocyte constitutes an invaluable system for studying cell polarity, the peculiar structure of its cytoskeleton prevents analysis at the level of microfilaments. Therefore the role of Moe in developing nurse cells was examined. At stage 9-10, the cellular face in contact with follicle cells presents a dense network of microfilaments that is organized in a mesh-like structure. Moesin accumulates strongly at the cortex of stage 9-10 nurse cells and colocalizes with this F-actin mesh. In DmoeX5 germline clones, the regular arrangement of the cortical actin network is lost. Instead, microfilaments are localized in a few aggregates that radiate from the cell surface, forming abnormal structures that resemble flowers. In DmoePL106 clones, F-actin is often absent from the cortex and concentrated at the cell perimeter. Although defects are more pronounced in the cortical region, F-actin also accumulates in abnormal structures at the point of contact between nurse cells. However, Moe mutations affect neither actin cables nor the actin-rich ring canals. These data show that Moe is specifically required in nurse cells for the organization of the cortical actin network, where Moesin strongly accumulates during stage 9-10. Therefore, this constitutes a suitable system to analyse the effects of Moesin Thr 559 substitutions on microfilament organization. In nurse cells expressing Moesin-TD, the mesh-like structure disappears and F-actin is concentrated in dense spots in the cortex and lateral membranes. Overall cell shape is altered and nurse cells become abnormally round. Expression of Moesin-TA also affects the organization of cortical microfilaments, which are enriched in 'actin-flowers' reminiscent of those observed in weak Moe mutants. Interestingly, Moesin-TA associates with the membrane, where it colocalizes with F-actin at the periphery of these aberrant structures. Individual microfilaments, which display a dotted-line staining characteristic of actin cables, are observed at the centre of the mutant structure. Altogether, these results provide the first genetic evidence that the regulated activity of Moe is required for the organization of a specific subset of microfilaments that are asymmetrically localized in the cell (Polesello, 2002).

Thus Moesin is required for the establishment of the antero-posterior body axis in Drosophila. Evidence is provided that Moesin is involved in actin-based functional cell asymmetry and morphogenesis during oogenesis. The divergence of Ezrin, Radixin and Moesin from a common ancestor is likely to be a relatively recent event that is restricted to deuterostomes. It may have been even more recent, as the Ciona intestinalis genome contains only one ERM gene. Therefore, Ezrin, Radixin and Moesin seem to be closely related paralogues, probably displaying a functional similarity that complicates their analysis during mammalian development. In contrast, the existence of the single ERM gene in Drosophila, Moesin, facilitates its functional analysis and might provide original clues for the understanding of ERM activity during development (Polesello, 2002).

Moesin is required in vivo for the proper anchorage of actin microfilaments to the PM. During oogenesis, mutations in Moe prevent the correct attachment of F-actin to the oocyte cortex. This has marked developmental consequences, as it correlates with the mislocalization of posterior determinants that disrupts the antero-posterior polarity of the future embryo. Several lines of evidence support the conclusion that the mislocalization of Oskar observed in Moe oocytes is a direct consequence of the disorganization of microfilaments. (1) Injection of microfilament-depolymerizing drugs into early embryos disrupts anchorage of the posterior determinants, and in common with the situation in Moe mutants, it does so without affecting formation of the anterior pole or altering dorso-ventral asymmetry. (2) Cytoplasmic Tropomyosin, a component of the actin cytoskeleton, is also required for accumulation of Oskar at the posterior end. (3) Drosophila Profilin (Chic) and Cap, two proteins involved in actin polymerization, are also required for the localization of Oskar and Staufen. However, the polarity defects caused by chic and cap mutations may be caused, at least in part, by disorganization of microtubules. In contrast, microtubule-dependent processes, such as localization of Gurken, nuclear migration and cytoplasmic streaming, are not affected in Moe mutants. Although the possibility that microtubules are altered in the total absence of Moe products cannot be excluded, both minus- and plus-end microtubule motors are properly localized in the Moe alleles analysed. Therefore, the abnormal distribution of posterior determinants observed in these Moe mutants is not a consequence of the disruption of microtubule nucleation or polarity. (4) The presence of Oskar protein in ectopic actin clumps, caused by mutations in Moe, further supports the hypothesis that alteration of the actin cytoskeleton is responsible for the mislocalization of Oskar. These data suggest a model in which after the microtubule-mediated transport of oskar mRNA to the posterior pole, Moesin is required to organize actin filaments and tether them to the membrane, where they then anchor oskar RNA and protein to the cortex (Polesello, 2002).

Drosophila Moesin also contributes to the cellular asymmetry of nurse cells, in which Moe is required for the organization of the cortical F-actin network. Interestingly, in weak Moe alleles, microfilaments make abnormal structures that radiate from the cell surface. This suggests that ERMs are required not only to anchor preformed microfilaments, but may also be required to assemble actin filaments close to the PM. Moreover, although ERMs are known to control protuberant actin-rich structures, this study reveals the determinant role of Moesin in the organization of a 'flat' polarized actin cytoskeleton. However, Moesin does not seem to be involved in general actin organization, since Moe mutations do not significantly affect other actin-dependent processes examined (Polesello, 2002).

To bind F-actin and membrane proteins, ERMs must first be activated by a mechanism that involves the phosphorylation of a conserved C-terminal threonine. Specific substitutions of this residue (Thr 559) affect the subcellular localization of Moesin in vivo. A phosphomimetic mutation redirects the Moesin protein to the membrane, where it colocalizes with F-actin. In agreement with ex vivo data obtained with mammalian ERM, this shows that the modification of Thr 559 triggers or stabilizes the interaction of Moesin with the membrane/cytoskeletal machinery in vivo. However, a mutant that prevents phosphorylation of Moesin on Thr 559 is still capable of associating to membranes, indicating that the membrane targeting of ERM also depends on other determinants, such as binding to phosphatidylinositol-4,5-bisphosphate and/or self-association. In addition, Moesin-TA associates with disorganized microfilaments at the nurse cell cortex, suggesting that phosphorylation of Thr 559 is not an absolute prerequisite for actin binding. Consistently, isolated Dmoesin C terminus binds ubiquitously to F-actin in vivo, even in tissues in which Moe is neither expressed nor required (and thus is unlikely to be properly regulated). In addition to membrane targeting, the dynamic regulation of Moesin activity through phosphorylation of Thr 559 is required for Moesin function during oogenesis. Whereas expression of Moesin-TA results in a scattered organization of interspersed microfilaments, expression of Moesin-TD causes the formation of very dense F-actin clumps and individual microfilaments are no longer observable. Interestingly, expression of Moesin-TA and Moesin-TD also has antagonistic effects on the distribution of Oskar, suggesting that the intrinsic organization of the actin cytoskeleton is actively involved in the proper localization of polarity determinants. The identification of the factors involved in the phosphorylation of Dmoesin Thr 559 now awaits the functional analysis of candidate protein kinases and the study of their activities with respect to Dmoesin (Polesello, 2002).

Moesin contributes an essential structural role in Drosophila photoreceptor morphogenesis

Ezrin-Radixin-Moesin (ERM) family proteins organize heterogeneous sub-plasma membrane protein scaffolds that shape membranes and their physiology. In Drosophila oocytes and imaginal discs, epithelial organization, fundamental to development and physiology, is devastated by the loss of Moesin. Moesin is crucial for Drosophila photoreceptor morphogenesis. Beyond its requirement for retinal epithelium integrity, Moesin is essential for the proper assembly of the apical membrane skeleton that builds the photosensitive membrane, the rhabdomere. Moesin localizes to the rhabdomere base, a dynamic locus of cytoskeletal reorganization and membrane traffic. Downregulation of Moesin through RNAi or genetic loss of function profoundly disrupts the membrane cytoskeleton and apical membrane organization. Normal levels and distribution of Moesin were found in photoreceptors of a Moesin mutant previously regarded as protein null, suggesting alternative interpretations for studies using this allele. These results show an essential structural role for Moesin in photoreceptor morphology (Karagiosis, 2004).

Confocal immunofluorescence was used to observe the membrane cytoskeleton during photoreceptor differentiation. Adult Drosophila photoreceptor apical plasma membranes are organized into complementary domains: the photosensory rhabdomere and an adjacent supporting membrane, the stalk. Rhabdomeres are columns of closely packed, photosensitive microvilli; a collar of stiffened membrane, the stalk, flanks rhabdomeres and supports them on the optical axis of the eye. Prominent coated pits and vesicles in the stalk pocket suggest a special role for the stalk in endocytosis. Molecular specialization of the apical membrane cytoskeleton supports rhabdomeres: microfilaments thread each microvillus, and a terminal web of F-actin extends from the rhabdomere base into the cytoplasm. Activated, phosphorylated Moesin (p-Moesin) localizes to the rhabdomere base and the polarity determinant transmembrane protein Crumbs localizes to the stalk (Izaddoost, 2002; Pellikka, 2002; Karagiosis, 2004 and references therein).

Definitive rhabdomere and stalk primordia are established within the photoreceptor apical membrane at approximately 50% of pupal development (% pd). Future rhabdomeres are covered with short, finger-like microvilli. Smooth membrane defines developing stalks of R2, R4 and R7, and the asymmetrical stalk of R5; R1, R3 and R6 lack stalks at this stage. Concomitant with morphological differentiation, Moesin and Crumbs resolve to their respective adult domains. The complementary distribution of these determinants of membrane/cytoskeleton interaction suggests that photoreceptor Moesin does not partner with Crumbs, as reported for embryonic epithelia (Medina, 2002). Before this stage, Moesin and Crumbs extend fully across the apical surface. The absence of stalks and the lack of Crumbs staining in photoreceptors R1, R3 and R6 suggests Moesin does not require Crumbs to establish the rhabdomere base (Karagiosis, 2004).

In order to visualize both active phosphorylated and dormant non-phosphorylated Moesin, Gal4/UAS-targeted expression of full-length wild-type Myc- and GFP-tagged Moesin was used. In fixed and live cells, both tagged proteins distribute throughout the cytoplasm and concentrate at the apical membrane. Cytoplasmic staining probably represents the soluble, dormant Moesin (Karagiosis, 2004).

To test whether the GFP-tagged Moesin is functionally active, the transgene was introduced into MoePL54 (Polesello, 2002), a homozygous lethal Moe- background. Constitutive expression of Moesin-GFP using the Act5C>Gal4 promoter rescues MoePL54 viability. Ommatidia examined with confocal microscopy and electron microscopy frequently exhibit rhabdomere size and organization defects, including microvillar gaps. Moesin dosage may play a crucial role in the proper organization of the apical surface (Karagiosis, 2004).

Since epithelial organization (which defines the context of photoreceptor differentiation), fails in flies lacking Moesin (Speck, 2003), IR-Moesin, a transgene carrying an inverted repeat sequence encoding a portion of Moesin was expressed to induce RNAi and downregulate Moesin late in eye development. Epithelial integrity is preserved in these eyes, permitting dissection of the role of Moesin in photoreceptor differentiation. The efficiency of UAS IR-Moesin downregulation of Moesin expression was assayed using immunolocalization. Using the heat shock>Gal4 driver (hs>Gal4) to drive the expression of UAS IR-Moesin in staged pupae, photoreceptor morphogenesis was found to be most sensitive to Moesin loss at the time of rhabdomere/stalk resolution, approximately 50% pd. At this stage, hs>IRMoesin expression completely abolished Moesin immunodetection in many photoreceptors. The membrane cytoskeleton, assayed by F-actin staining, is disorganized in photoreceptors lacking Moesin. Rhabdomeres of these eyes typically have reduced, irregular microvillar fields (Karagiosis, 2004).

Expression of UAS IR-Moesin using the GMR>Gal4 driver, which becomes active soon after photoreceptors are fated in the retinal epithelium, severely disrupts actin cytoskeleton organization and apical differentiation. Distinct microvillar and stalk membranes are apparent, but are discontinuous and jumbled. It remains to be determined whether higher levels of IR-Moesin, or its expression during an especially sensitive period, account for the stronger phenotype of GMR>IR-Moesin eyes. Downregulation of Moesin during photoreceptor differentiation focuses defects on the rhabdomere; cells appear otherwise healthy and possess apparently normal adherens junctions, probably accounting for the preservation of epithelial integrity. Once established, adherens junctions may not require Moesin to maintain epithelial integrity; alternatively, severely reduced levels may be sufficient (Karagiosis, 2004).

As an alternate strategy to remove zygotic Moesin later in development, eye clones were generated homozygous for either of two Moesin alleles, MoePL54 and MoeX5, shown to be null or strong hypomorphs (Polesello, 2002). In pupal MoePL54 mosaic eyes, containing a mixture of Moesin-positive cells and cells with severely reduced Moesin, rhabdomeres of photoreceptors with immuno-undetectable Moesin show severely disrupted apical membranes. Assayed with rhodamine-phalloidin, the rhabdomere microvillar array in Moe- cells is irregular and the RTW is replaced with abnormal F-actin accumulations. In MoeX5 mosaic eyes, cells lacking Moesin are rarely observed, but ommatidia containing reduced numbers of photoreceptors are common, suggesting that the missing cells died or were otherwise lost from the developing retinal epithelium (Karagiosis, 2004).

Notably, a third hypomorphic allele, MoeG0323, in a Rho-reduced background, which lacks Moesin during larval development (Speck, 2003), contains normal levels of Moesin at the rhabdomere base. Both hemizygous MoeG0323/Y; Rho172R/CyO and homozygous MoeG0323/MoeG0323; Rho172R/CyO adult retinas show normal photoreceptors and Moesin localization to the rhabdomere base. Western analysis of whole head extracts of these flies shows that Moesin is present, but at a reduced level (Karagiosis, 2004).

To examine the role of Moesin activation in photoreceptor morphogenesis, a transgenic line was generated expressing a constitutively active phosphomimetic mutation, UAS T559D Moesin Myc, under Gal4/UAS control. When hs>T559D Moesin Myc is expressed during apical morphogenesis, a profusion of irregular microvilli dominates the apical surface. This result parallels observations in cell culture, where expression of T559D Moesin results in hyper-formation of irregular microvilli. Other cellular structures, including the adherens junctions, appear to be intact. Using confocal immunofluorescence, T559D Moesin Myc localizes to the entire photoreceptor plasma membrane and concentrates at the apical membrane. The failure of basolateral Moesin to generate microvilli suggests that additional, apically-limited proteins are needed to initiate rhabdomere microvilli. T559D Moesin severely disrupts the actin cytoskeleton, and Crumbs, which should resolve to the stalk, remains distributed across the entire apical surface. Failure of Crumbs to re-localize is a harbinger of the loss of stalk/rhabdomere distinction in photoreceptors expressing dominant-active Moesin. It is speculated that dynamic turnover and regulation of the protein is important for morphogenesis, and the unregulated binding of dominant-active Moesin 'freezes' the apical membrane, compromising its reorganization (Karagiosis, 2004).

Epithelial cell apical plasma membranes host elaborate membrane and sub-membrane protein assemblies that support sensory and effector mechanisms and are often amplified and highly structured by the actin membrane cytoskeleton. Substantial in vitro and in vivo evidence implicates Moesin in the genesis and regulation of this membrane cytoskeleton, and these results extend this connection to a role for Moesin in the morphogenesis of a complex, compartmented apical specialization, the Drosophila rhabdomere. The dynamic distribution of Moesin during photoreceptor development and the impact of Moesin gain-and loss-of-function during morphogenesis suggest an essential role for the protein in organizing the membrane skeleton that supports the microvillar array of the rhabdomere (Karagiosis, 2004).

The developmental cues that restrict Moesin to the rhabdomere primordium at the onset of overt morphogenesis are not known. At this stage, photoreceptor apices contact each other in a stereotyped pattern of apical cell-cell contacts within a trapped apical pocket that will later open to form the IRS, suggesting that apical contacts may localize future rhabdomeres. In other systems, ERM proteins are recruited to plasma membranes and activated there by PtdIns(4,5)P2-binding and phosphorylation of a C-terminal threonine (Hirao, 1996; Matsui, 1998). Rhabdomeres are rich in PtdIns(4,5)P2, and Rhodopsin-activated cleavage of PtdIns(4,5)P2 by the phospholipase C. NorpA is the substrate for Drosophila phototransduction. A Moesin-organized nexus of phosphoinositide and actin organizing proteins at the rhabdomere base may represent an economical dual use of the system for structure and physiology (Karagiosis, 2004).

The restriction of Moesin to the rhabdomere base and its loss-of-function phenotypes suggests that Moesin links microvillar cytoplasmic ends to the underlying actin cytoskeleton, the rhabdomere terminal web (RTW). The full molecular makeup of the Moesin-organized photoreceptor cytoskeleton and the nature of the forces it organizes during morphogenesis remain to be determined, but the catenary-like deformation of the rhabdomere base as developing microvilli elongate suggests the membrane cytoskeleton contributes a sub-apical constraint, a tensile sheet that contains the expanding photosensitive membrane in an orderly microvillar stack. In other systems, Moesin-organized protein scaffolds recruit potent regulators of membrane architecture and function, such as the small GTPase regulators RhoGDI, RabGAPs and RhoGAPs. It is plausible that the ability of Moesin to bind F-actin and its regulators may contribute to the establishment of the RTW. Rhabdomere membrane is in dynamic exchange across the rhabdomere base with endosomes of the photoreceptor cytoplasm, and it is notable that ERM proteins bind to regulators of membrane recycling. Moesin may participate in the membrane exchange that supports a dynamic sensory membrane (Karagiosis, 2004).

Recent observation that the lethality of flies homozygous for MoeG0323, considered a protein null, could be rescued in Rho-reduced flies was interpreted as showing that Moesin facilitates epithelial morphology not by providing an essential structural function, but rather by antagonizing activity of the small GTPase Rho (Speck, 2003). This conclusion rests on MoeG0323 being a protein null. The observations that Rho-reduced MoeG0323 produces Moesin detectable by Western analysis and shows photoreceptor Moesin immunostaining indistinguishable from wild type suggest alternate interpretations. One possibility is that the P-element insertion in the 5' untranslated region of MoeG0323 downregulates embryonic and larval Moesin, but can be over-ridden later in development in Rho-reduced animals. Such interpretation, while not ruling out a role for Moesin in Rho regulation, is compatible with present and previous evidence of a structural role for Moesin (Karagiosis, 2004).

Given the fundamental parallels between Drosophila and vertebrate photoreceptors, it is interesting to consider whether the ERM-organized membrane cytoskeleton will play a similar role in human photoreceptor development and disease (Karagiosis, 2004).

Slik Sterile-20 kinase regulates Moesin activity to promote epithelial integrity during tissue growth

The Drosophila Sterile-20 kinase Slik (FlyBase name: Polo kinase kinase 1) promotes tissue growth during development by stimulating cell proliferation and by preventing apoptosis. Proliferation within an epithelial sheet requires dynamic control of cellular architecture. Epithelial integrity fails in slik mutant imaginal discs. Cells leave the epithelium and undergo apoptosis. The abnormal behavior of slik mutant cells is due to failure to phosphorylate and activate Moesin, which leads to excess Rho1 activity. This is distinct from Slik's effects on cell proliferation; these effects are mediated by Raf. Thus Slik acts via distinct pathways to coordinate cell proliferation with epithelial cell behavior during tissue growth (Hipfner, 2004).

The wing imaginal disc is an epithelial sac composed of two distinct but continuous epithelial layers enclosing a central lumen. The portion of the disc that will form the wing is a pseudostratified columnar epithelium. In optical cross-section, the dense packing of the cells is visible, with nuclei appearing stacked in layers. Overlying the columnar epithelium is the squamous peripodial epithelium. The apical surface of both epithelial layers is oriented toward the lumen of the disc, as seen by the concentration of F-actin near the adherens junctions. In wing discs lacking Slik activity, the columnar epithelium is abnormally thin. Many cells lost their capacity to remain integrated in the epithelium and are extruded basally to form a disorganized mass. Many of these cells undergo apoptosis as evidenced by pyknotic nuclei and by TUNEL labeling, though clusters of cells survived. A similar but milder defect was seen in discs with reduced Slik activity. Many apoptotic cells with activated Caspase are seen in optical sections below the epithelial layer. Some of the extruded slik mutant cells are alive and appeared mesenchymal, having lost their polarized epithelial character. These cells produce F-actin-rich filopodia and appear to acquire motile behavior. Thus slik activity helps cells to maintain epithelial integrity (Hipfner, 2004).

The observation that many live slik mutant cells are extruded from the epithelium suggests that loss of epithelial integrity is not a consequence of apoptosis. This was confirmed by producing clones of slik mutant cells that expressed the baculovirus caspase inhibitor p35. p35-expressing slik mutant cells also lose epithelial integrity and are extruded from the epithelium, but remain alive. This suggests that apoptosis is a consequence of the loss of epithelial organization, perhaps due to loss of survival signaling by Raf (Hipfner 2004; Hipfner, 2004).

To investigate whether slik is essential for epithelial polarity per se, the subcellular localization of junctional complex proteins was compared in normal cells and clones of slik1 null mutant cells. In wild-type cells, Slik protein is concentrated apically and colocalizes with cortical actin, in addition to a diffuse cytoplasmic staining. Slik is concentrated apical to septate junctions (marked by Discs large) and adherens junctions (marked by E-Cadherin and anti-phosphotyrosine) and to the apical-most marginal zone complexes containing the PDZ-domain protein Patj, indicating that Slik is at or near the apical membrane rather than in the junctions. Junctional complexes appear normal in slik mutant cells that remain in the disc epithelium, but are lacking in mutant cells that have left the epithelium. Thus, although it is not essential for apical-basal polarity, slik contributes to maintaining epithelial organization. Cells lacking slik often lose epithelial polarity and leave the epithelium. Many of these cells undergo apoptosis (Hipfner, 2004).

slik1 mutant clones generated in the eye disc early in development and provided with a growth advantage can grow to large sizes. As in the wing disc, many cells are extruded from the eye disc and undergo apoptosis, whereas others remain integrated in the epithelium and show normal expression of the neuronal marker ELAV in photoreceptors. Adult eyes with large slik1 mutant clones show moderate external roughness. Although many properly organized ommatidial clusters in these eyes are found, mutant photoreceptors show defects during subsequent differentiation. Each photoreceptor normally projects a stack of actin-based microvilli, or rhabdomere, from the apical membrane domain that provides the increased membrane surface for harvesting light. Rhabdomeres form during pupal development (PD). By 70% PD, the eight photoreceptors that comprise a single facet in the wild-type eye have begun to form rhabdomeres consisting of short bundles of microvilli. The cells are connected to each other by adherens junctions, which separate the apical from the basolateral membrane domains. At 95% PD the microvilli have extended to form regular bundles. The arrangement of photoreceptors is largely normal in complete ommatidia containing slik mutant clones, but many slik mutant photoreceptors had patches of apical membrane devoid of microvilli. Where present, the microvilli in slik mutant photoreceptors do not form organized stacks. Mutant photoreceptors are present in eyes from 18-day-old adult flies, indicating that their survival as postmitotic cells is not impaired. These defects resemble those described recently (Karagiosis, 2004) in photoreceptors lacking Moesin (Hipfner, 2004).

The similarity in slik and moesin mutant phenotypes prompted a comparison of the defects in imaginal discs in more detail. Loss of Moesin causes defects in epithelial integrity due to Rho1-induced changes in the actin cytoskeleton (Speck, 2003). Interestingly, cells extruded from moesin mutant discs also undergo massive apoptosis. The organizational changes in the actin cytoskeleton are strikingly similar in the two genotypes. These defects in moesin mutant discs can be rescued by reduction of rho1 gene dosage (Speck, 2003). Similarly, the defects in epithelial integrity and cell survival in hypomorphic slik mutants are strongly suppressed when rho1 gene dosage is reduced, suggesting that some defects in slik mutants are due to excessive Rho1 activity and that Slik acts in the same pathway as Moesin (Hipfner, 2004).

Moesin is activated by phosphorylation on a conserved Threonine residue in the C-terminal domain. Thr 556-phosphorylated Moesin (P-Moesin) colocalizes with apical Slik protein in wild-type cells, in addition to a diffuse distribution of both proteins. P-Moesin levels are considerably reduced in slik1 clones, compared to adjacent Slik-expressing cells in the wing disc. Immunoblot analysis showed that Thr 556-phosphorylation is reduced in slik mutant wing discs; the level of total Moesin is unchanged. Likewise, Slik RNAi in S2 cells results in a dose-dependent decrease in endogenous Slik protein and in Moesin phosphorylation. Expression of the kinase domain of Slik is sufficient to induce PMoesin in the wing disc, whereas a catalytically inactive version of this domain has no effect. Activated Moesin has been shown to stabilize F-actin (Speck, 2003), and increased F-actin staining is observed in Slik kinase domain-expressing cells. These data indicate that Slik activity is required for Moesin Thr 556-phosphorylation in vivo. However, it was not possible to direct phosphorylation of Moesin by Slik in vitro, suggesting that Slik may act indirectly to promote Moesin phosphorylation in vivo (Hipfner, 2004).

Speck (2003) showed that activated Moesin limits Rho activity in vivo. Experiments in cultured mammalian cells have suggested that Rho might also act upstream of Moesin to promote its phosphorylation, forming a feedback loop (Matsui, 1998; Shaw, 1998; Tran Quang, 2000). To assess the involvement of Rho in Moesin phosphorylation, the effects of removing Slik, Rho1, and the Rho effector kinase Rok from S2 cells were compared by RNAi. Depletion of endogenous Slik strongly reduces Moesin phosphorylation. Most cells depleted of Slik show low levels of Moesin phosphorylation. Expression of the Slik kinase domain in the Slik-depleted cells restores Moesin phosphorylation to an extent that is consistent with the transfection efficiency in the experiment. Depletion of Rho1 causes a modest reduction in Moesin phosphorylation, and expression of the Slik kinase domain increases the level of Moesin phosphorylation in Rho1-depleted cells. Many cells depleted of Rho1 show levels of Moesin phosphorylation similar to those of control S2 cells, and cells expressing the Slik kinase domain are always among those with the higher level of Moesin phosphorylation. This indicates that Rho1 is not required for Slik-induced phosphorylation of Moesin in S2 cells. The mammalian kinase Rock has been suggested to serve as an effector of Rho in mediating ERM protein phosphorylation, though this is controversial. In this study, depletion of the Drosophila ortholog, Rok, from S2 cells had little or no effect on the level of Moesin phosphorylation, and did not prevent Slik kinase domain-induced Moesin phosphorylation. These observations suggest that Rho1 is not required for Moesin phosphorylation in S2 cells. Although the level of Rho activity is elevated in slik mutants, the level of Moesin phosphorylation is very low, suggesting that Rho activity cannot compensate for the lack of Slik activity (Hipfner, 2004).

Next it was asked whether the epithelial defects in slik mutants could be rescued by expression of a constitutively active, phosphomimetic form of Moesin (MoesinTD; Speck, 2003). moeGO323 mutants served as a control. Moesin protein was nearly undetectable in moeGO323 discs by immunoblot. In moeGO323 discs, armGAL4 drove ubiquitous MoesinTD expression to a level lower than endogenous Moesin. This level of activated Moesin had no effect on the development of wild-type animals. moeGO323 mutants grew more slowly than wild-type larvae, reached pupal stages later, and died as pupae. More than half of the mutant larvae were rescued to pharate adult (25%) or adult stages (28%) by expression of MoesinTD (vs. 0% and 1% for moeGO323). MoesinTD expression also partially rescued slik1 mutants. Nearly half of the slik1 animals survived to pupation, but all died shortly thereafter. In contrast, 9% of slik1 mutants expressing MoesinTD reached the pharate adult stage. One small rescued slik1 mutant fly survived to adulthood. Discs from rescued slik1 mutants showed some improvement in overall structure and actin organization. Basal extrusion of cells was reduced and in some cases nearly eliminated (Hipfner, 2004).

MoesinTD expression did not rescue slik1 animals to the same extent as moeGO323 mutants. Despite improvements in epithelial integrity and the stage of lethality, MoesinTD had little or no effect on the slow rate of development and growth of slik1 mutants. One explanation for this may be that Moesin acts downstream of Slik to regulate epithelial integrity but not to promote growth. Thus the requirement for Moesin in Slik-driven tissue growth was examined. The effect of Slik on cell proliferation is easily observed in the peripodial cells overlying the columnar wing disc epithelium. Expression of Slik in the columnar epithelium induces nonautonomous proliferation of peripodial cells, detectable as an increase in nuclear density and BrdU incorporation. The same effect was observed when Slik was expressed in moeGO323 mutants. Activated MoesinTD does not affect cell proliferation. Thus Moesin is not necessary for Slik-induced cell proliferation, nor is Moesin activity sufficient to stimulate cell proliferation (Hipfner, 2004).

It is concluded that Slik regulates epithelial integrity via Moesin activation, independent of its effects on tissue growth. Slik-induced tissue growth is Raf-dependent. Expression of a kinase inactive form of Slik is capable of promoting Raf-dependent overproliferation, suggesting that the growth effect may be mediated by protein-protein interactions rather than by Slik kinase activity (Hipfner, 2003). Consistent with this notion, expression of the kinase domain of Slik alone does not induce cell proliferation or tissue overgrowth. In contrast, Moesin phosphorylation is dependent on Slik kinase activity. Taken together, these observations suggest two distinct effector pathways for Slik -- a pathway controlling growth, involving Raf, and a separate pathway controlling epithelial integrity involving Moesin phosphorylation (Hipfner, 2004).

What might be the purpose of the dual activities of Slik? Slik is not essential for cells to grow and divide, but it does control the rate of cell proliferation. Slik activity must be maintained within a defined range; too much or too little activity results in apoptosis (Hipfner, 2003). Mitogenic signaling through Slik can promote tissue growth and at the same time reinforce cellular architecture by maintaining Moesin in an active state. Proliferation-induced apoptosis prevents excessive Slik signaling from deregulating proliferation, as has been suggested for certain oncogenes. Under conditions of reduced mitogenic signaling, decreased Slik activity would result in a lower rate of cell proliferation and, by reducing Moesin activity, make cells more likely to be extruded from the disc and thus to undergo apoptosis. A more direct function of ERM proteins in regulating apoptosis may also be involved (Gautreau, 1999; Parlato, 2000). Thus, Slik activity may serve as a switch between pro-proliferative and pro-apoptotic states. Consistent with this idea, loss of ERM protein phosphorylation and activity has been shown to be an early event in apoptosis (Kondo, 1997). Interestingly, it is known that slowly dividing cells in Drosophila imaginal discs undergo apoptosis as a result of reduced ability to compete for survival factors. The dual activity of Slik may help to ensure elimination of less fit cells (Hipfner, 2004).

Rho GTPase controls Drosophila salivary gland lumen size through regulation of the actin cytoskeleton and Moesin

Generation and maintenance of proper lumen size is important for tubular organ function. This study reports on a novel role for the Drosophila Rho1 GTPase in control of salivary gland lumen size through regulation of cell rearrangement, apical domain elongation and cell shape change. Rho1 controls cell rearrangement and apical domain elongation by promoting actin polymerization and regulating F-actin distribution at the apical and basolateral membranes through Rho kinase. Loss of Rho1 results in reduction of F-actin at the basolateral membrane and enrichment of apical F-actin, the latter accompanied by enrichment of apical phosphorylated Moesin. Reducing cofilin levels in Rho1 mutant salivary gland cells restores proper distribution of F-actin and phosphorylated Moesin and rescues the cell rearrangement and apical domain elongation defects of Rho1 mutant glands. In support of a role for Rho1-dependent actin polymerization in regulation of gland lumen size, loss of profilin (Chickadee) phenocopies the Rho1 lumen size defects to a large extent. Ribbon, a BTB domain-containing transcription factor, functions with Rho1 in limiting apical phosphorylated Moesin for apical domain elongation. These studies reveal a novel mechanism for controlling salivary gland lumen size, namely through Rho1-dependent actin polymerization and distribution and downregulation of apical phosphorylated Moesin (Xu, 2011).

Rho1 acts both in salivary gland cells and in the surrounding mesoderm to maintain apical polarity during gland invagination and to mediate cell shape change during gland migration. This study demonstrates a novel role for Rho1 in controlling salivary gland lumen size through regulation of actin polymerization and distribution and regulation of Moesin activity. By analyzing Rho1 alleles for which salivary gland cells invaginated and formed a gland, it was shown that zygotic loss of function of Rho1 results in shortening and widening of the gland lumen, which is accompanied by defects in cell shape change and cell rearrangement and failure of apical domains to elongate along the Pr-Di axis of the gland. These effects of Rho1 are mediated through Rok; inhibition of Rok completely phenocopies loss of Rho1 in these cellular events. Based on these studies, a model is proposed for Rho1 control of salivary gland lumen size, in particular lumen width, which is determined by cell rearrangement and apical domain elongation. Rho1 and Rok, through inhibition of cofilin, regulate cell rearrangement and apical domain elongation by promoting actin polymerization to localize F-actin at the basolateral membrane and by limiting the apical accumulation of F-actin. In parallel to its role in actin polymerization and distribution, Rho1 acts independently of Rok to limit apical p-Moe with Rib by an unknown mechanism and this function of Rho1 is specific for apical domain elongation. The data on cofilin (Twinstar) are consistent with those in cultured HeLa cells that showed that mammalian ROCK can inhibit cofilin activity indirectly through LIMK-mediated phosphorylation of cofilin (Xu, 2011).

Although manipulating Moe activity through gland-specific expression of MoeT559D was sufficient to completely phenocopy the Rho1 lumen defects, including cell rearrangement, it did so without disrupting actin polymerization or distribution. This is likely to be due to activated Moe strengthening the link between the actin cytoskeleton and the apical plasma membrane (without affecting levels of apical F-actin), which would increase apical membrane stiffness and remove the ability of gland cells to rearrange. Indeed, Moesin has been shown to control cortical rigidity during mitosis of cultured Drosophila S2R+ cells. Thus, Rho1 regulates cell rearrangement and apical domain elongation by controlling the actin cytoskeleton and Moesin activity through distinct mechanisms (Xu, 2011).

The observation that chic mutant glands phenocopy Rho1 mutant glands to a large extent, suggests that Rho1 control of salivary gland lumen size is mainly dependent on a requirement for Rho1 in actin polymerization. However, as the chic and Rho1 gland lumen phenotypes are not identical, with chic mutant glands lacking the apical accumulation of F-actin and p-Moe observed in Rho1 mutant glands, Rho1 probably has an additional function in limiting accumulation of F-actin and p-Moe at the apical membrane. This function of Rho1, at least for limiting apical F-actin, might partly involve Rab5- or Shi-mediated endosome trafficking, because inhibition of Rab5 alone or Shi alone led to accumulation of F-actin at the apical membrane. Although Rab5DN- or ShiDN-expressing salivary gland cells were enriched with apical F-actin, lumen size was not affected. This could be due to Rab5DN and ShiDN affecting a pool of apical F-actin distinct from that affected by Rho1 and/or because Rab5DN-expressing gland cells retain basolateral F-actin and the ratio of apical to basolateral F-actin is not altered sufficiently to cause lumen size defects. In Rho11B mutant gland cells, some early endosomes were not coated with F-actin. Actin is known to contribute to multiple steps of the endocytic pathway, including movement of endocytic vesicles through the cytoplasm and their transport to late endosomes and lysosomes. One possible mechanism by which Rho1 normally limits apical accumulation of F-actin is by promoting its removal from the apical membrane and accumulation on endocytic vesicles (Xu, 2011).

Currently, it is not know how Rho1 limits accumulation of apical p-Moe. Membrane localization and activity of Moesin can be regulated via a number of mechanisms, such as its phosphorylation on a conserved Threonine residue, binding to phosphatidylinositol-(4,5)bisphosphate [PtdIns(4,5)P2] and association with components of the sub-membrane cytoskeleton, such as Crb. Studies in cultured mammalian cells have demonstrated that Rho signaling activates Moe either through phosphorylation of Moe by ROCK or through ROCK-mediated inhibition of myosin phosphatase, which is known to dephosphorylate p-Moe. Although it is possible that Drosophila Rho1 positively regulates Moe activity by one or more of these mechanisms, this study shows that in the developing salivary glands Rho1 in fact negatively regulates Moe activity. In rib mutant embryos, in which p-Moe is enriched apically, salivary gland and tracheal cells showed decreased staining for Rab11 GTPase, which localizes to the apical recycling endosomes and to secretory vesicles destined for the apical membrane. Thus, Rho1, like Rib might limit apical p-Moe through its membrane transport (Xu, 2011).

In Drosophila imaginal disc epithelia, Moe negatively regulates Rho1 activity to maintain epithelial integrity and to promote cell survival. These studies demonstrating that in the developing salivary gland Rho1 antagonizes Moe activity by limiting its localization at the apical membrane, shed novel insight into the functional relationship between Rho1 and Moe. It is possible that in a dynamic epithelium, such as the developing salivary gland, Rho1 contributes to the precise spatial and temporal regulation of Moe activity to fine-tune selective changes in apical domain shape. By contrast, in the imaginal disc epithelium, Rho1 regulation of Moe might not be necessary and, instead, Moe regulation of Rho1 activity is required to maintain epithelial integrity and cell survival. Thus, Rho and Moe can antagonize each other's activities depending on the type of epithelia or cellular event (Xu, 2011).

Rescue studies with Rho1WT demonstrate that Rho1 functions predominantly in the salivary gland cells to control apical domain elongation and cell rearrangement. Interestingly, expression of Rho1WT in the mesoderm with twi-GAL4 has no effect on cell rearrangement and has little effect on apical domain elongation and lumen size, whereas it has been shown that Rho1WT expression in the mesoderm significantly rescues the gland migration defect of Rho11B mutant embryos. This suggests that gland migration and lumen size control are regulated by distinct mechanisms. In support of this conclusion, embryos mutant for multiple edematous wings, encoding the βPS1 integrin subunit, which have defects in gland migration, show no defects in gland lumen width. Identifying the distinct and overlapping mechanisms by which salivary gland lumen width and length are controlled will help to elucidate the mechanisms by which lumen size is controlled in tubular organs (Xu, 2011).

Seamless tube shape is constrained by endocytosis-dependent regulation of active moesin
Most tubes have seams (intercellular or autocellular junctions that seal membranes together into a tube), but 'seamless' tubes also exist. In Drosophila, stellate-shaped tracheal terminal cells make seamless tubes, with single branches running through each of dozens of cellular extensions. Mutations in braided impair terminal cell branching and cause formation of seamless tube cysts. This study shows that braided encodes Syntaxin7 and that cysts also form in cells deficient for other genes required either for membrane scission (shibire) or for early endosome formation (Rab5, Vps45, and Rabenosyn-5). These data define a requirement for early endocytosis in shaping seamless tube lumens. Importantly, apical proteins Crumbs and phospho-Moesin accumulate to aberrantly high levels in braided terminal cells. Overexpression of either Crumbs or phosphomimetic Moesin induced lumenal cysts and decreased terminal branching. Conversely, the braided seamless tube cyst phenotype was suppressed by mutations in crumbs or Moesin. Indeed, mutations in Moesin dominantly suppressed seamless tube cyst formation and restored terminal branching. It is proposed that early endocytosis maintains normal steady-state levels of Crumbs, which recruits apical phosphorylated (active) Moe, which in turn regulates seamless tube shape through modulation of cortical actin filaments (Schottenfeld-Roames, 2014).


Amieva, M. R., Litman, P., Huang, L. Q., Ichimaru, E. and Furthmayr, H. (1999). Disruption of dynamic cell surface architecture of NIH3T3 fibroblasts by the N-terminal domains of moesin and ezrin: in vivo imaging with GFP fusion proteins. J. Cell Sci. 112: 111-125. 9841908

Batchelor, C. L., Woodward, A. M. and Crouch, D. H. (2004), Nuclear ERM (ezrin, radixin, moesin) proteins: regulation by cell density and nuclear import. Exp Cell Res. 296(2): 208-22. 15149851

Baumgartner, M., et al. (2006). The Nck-interacting kinase phosphorylates ERM proteins for formation of lamellipodium by growth factors. Proc. Natl. Acad. Sci. 103(36): 13391-6. Medline abstract: 16938849

Brumby, A. M., Goulding, K. R., Schlosser, T., Loi, S., Galea, R., Khoo, P., Bolden, J. E., Aigaki, T., Humbert, P. O. and Richardson, H. E. (2011). Identification of novel Ras-cooperating oncogenes in Drosophila melanogaster: a RhoGEF/Rho-family/JNK pathway is a central driver of tumorigenesis. Genetics 188: 105-125. PubMed ID: 21368274

Carreno, S., et al. (2008). Moesin and its activating kinase Slik are required for cortical stability and microtubule organization in mitotic cells. J. Cell Biol. 180: 739-746. PubMed Citation: 18283112

Chorna-Ornan, I., et al. (2005). Light-regulated interaction of Moe with Trp and Trpl channels is required for maintenance of photoreceptors. J. Cell Biol. 171(1): 143-52. 16216927

Deretic, D., Traverso, V., Parkins, N., Jackson, F., Rodriguez de Turco, E. B. and Ransom. N. (2004). Phosphoinositides, ezrin/moesin, and rac1 regulate fusion of rhodopsin transport carriers in retinal photoreceptors. Mol. Biol. Cell 15: 359-370. 13679519

Doi, Y., Itoh, M., Yonemura, S., Ishihara, S., Takano, H., Noda, T. and Tsukita, S. (1999). Normal development of mice and unimpaired cell adhesion cell motility actin-based cytoskeleton without compensatory upregulation of ezrin or radixin in moesin gene knockout. J. Biol. Chem. 274: 2315-2321. 9890997

Drees, F., Pokutta, S., Yamada, S., Nelson, W. J., Weis, W. I. (2005). alpha-Catenin is a molecular switch that binds E-cadherinß-catenin and regulates actin-filament assembly. Cell 123: 903-915. 16325583

Edwards, K. A., et al. (1994). Identification of Drosophila cytoskeletal proteins by induction of abnormal cell shape in fission yeast. Proc. Natl. Acad. Sci. 91: 4589-4593. 8183953

Edwards, K. A., Demsky, M., Montague, R. A., Weymouth, N. and Kiehart, D. P. (1997). GFP-moesin illuminates actin cytoskeleton dynamics in living tissue and demonstrates cell shape changes during morphogenesis in Drosophila. Dev. Biol. 191(1): 103-17. 9356175

Faure, S., et al. (2004). ERM proteins regulate cytoskeleton relaxation promoting T cell-APC conjugation. Nat Immunol. 5(3):272-9. 14758359

Finnerty, C. M., et al. (2004). The EBP50-moesin interaction involves a binding site regulated by direct masking on the FERM domain. J. Cell Sci. 117(Pt 8) :1547-52. 1502068

Gatto, C. L., Walker, B. J. and Lambert S. (2003). Local ERM activation and dynamic growth cones at Schwann cell tips implicated in efficient formation of nodes of Ranvier. J. Cell Biol. 162(3): 489-98. 12900397

Gautreau, A., Poullet, P., Louvard, D. and Arpin, M. (1999). Ezrin, a plasma membrane-microfilament linker, signals cell survival through the phosphatidylinositol 3-kinase/Akt pathway. Proc. Natl. Acad. Sci. 96: 7300-7305. 10377409

Gautreau, A., Louvard, D. and Arpin, M. (2000). Morphogenic effects of ezrin require a phosphorylation-induced transition from oligomers to monomers at the plasma membrane. J. Cell Biol. 150: 193-203. 10893267

Hamada, K., et al. (2003). Structural basis of adhesion-molecule recognition by ERM proteins revealed by the crystal structure of the radixin-ICAM-2 complex. EMBO J. 22(3): 502-14. 12554651

Hayashi, K., Yonemura, S., Matsui, T. and Tsukita, S. (1999). Immunofluorescence detection of ezrin/radixin/moesin (ERM) proteins with their carboxyl-terminal threonine phosphorylated in cultured cells and tissues. J. Cell Sci. 112: 1149-1158. 10085250

Hegan, P. S., Mermall, V., Tilney, L. G. and Mooseker, M. S. (2007). Roles for Drosophila melanogaster Myosin IB in maintenance of enterocyte brush-border structure and resistance to the bacterial pathogen Pseudomonas entomophila. Mol. Biol. Cell 18(11): 4625-36. PubMed Citation: 17855510

Hipfner, D. R. and Cohen, S. M. (2003). The Drosophila Sterile-20 kinase Slik controls cell proliferation and apoptosis during imaginal disc development. PLoS. Biol. 1: 244-256. 14624240

Hipfner, D. R., Keller, N. and Cohen, S. M. (2004). Slik Sterile-20 kinase regulates Moesin activity to promote epithelial integrity during tissue growth. Genes Dev. 18(18): 2243-8. 15371338

Hirao, M., Sato, N., Kondo, T., Yonemura, S., Monden, M., Sasaki, T., Takai, Y. and Tsukita, S. (1996). Regulation mechanism of ERM (ezrin/radixin/moesin) protein/plasma membrane association: possible involvement of phosphatidylinositol turnover and Rho-dependent signaling pathway. J. Cell Biol. 135: 37-51. 8858161

Hsieh, H. H., Chang, W. T., Yu, L. and Rao, Y. (2014). Control of axon-axon attraction by Semaphorin reverse signaling. Proc Natl Acad Sci U S A. 111(31):11383-8. PubMed ID: 25049408

Hughes, S. C., Formstecher, E. and Fehon, R. G. (2010). Sip1, the Drosophila orthologue of EBP50/NHERF1, functions with the sterile 20 family kinase Slik to regulate Moesin activity. J. Cell Sci. 123: 1099-107. PubMed Citation: 20215404

Ivetic, A., Florey, O., Deka, J., Haskard, D. O., Ager, A. and Ridley, A. J. (2004) Mutagenesis of the ezrin-radixin-moesin binding domain of L-selectin tail affects shedding, microvillar positioning, and leukocyte tethering. J. Biol. Chem. 279(32): 33263-72. 1517869

Jankovics, F., Sinka, R., Lukacsovich, T. and Erdelyi, M. (2002). MOESIN crosslinks actin and cell membrane in Drosophila oocytes and is required for OSKAR anchoring. Curr. Biol. 12(23):2060-5. 12477397

Karagiosis, S. A. and Ready, D. F. (2004). Moesin contributes an essential structural role in Drosophila photoreceptor morphogenesis. Development 131(4): 725-32. 14724125

Kondo, T., Takeuchi, K., Doi, Y., Yonemura, S., Nagata, S. and Tsukita, S. (1997). ERM (ezrin/radixin/moesin)-based molecular mechanism of microvillar breakdown at an early stage of apoptosis. J. Cell Biol. 139: 749-758. 9348291

Kunda, P., Pelling, A. E., Liu, T. and Baum, B. (2008). Moesin controls cortical rigidity, cell rounding, and spindle morphogenesis during mitosis. Curr. Biol. 18: 91-101. PubMed Citation: 18207738

Lozupone, F., et al. (2004). Identification and relevance of the CD95-binding domain in the N-terminal region of ezrin. J. Biol. Chem. 279(10): 9199-207. 14676203

Mackay, D. J., Esch, F., Furthmayr, H. and Hall, A. (1997). Rho- and rac-dependent assembly of focal adhesion complexes and actin filaments in permeabilized fibroblasts: an essential role for ezrin/radixin/moesin proteins. J. Cell Biol. 138: 927-938. 9265657

Maeda, M., Doi, Y., Yonemura, S., Amano, M., Kaibuchi, K. and Tsukita, S. (1998). Rho-kinase phosphorylates COOH-terminal threonines of ezrin/radixin/moesin (ERM) proteins and regulates their head-to-tail association. J. Cell Biol. 140: 647-657. 9456324

Matsui, T., et al. (1998). Rho-kinase phosphorylates COOH-terminal threonines of ezrin/radixin/moesin (ERM) proteins and regulates their head-to-tail association. J. Cell Biol. 140: 647-657. 9456324

McCartney, B. M. and Fehon, R. G. (1996). Distinct cellular and subcellular patterns of expression imply distinct functions for the Drosophila homologues of moesin and the neurofibromatosis 2 tumor suppressor, merlin. J. Cell Biol. 133: 843-852. 8666669

Medina, E., et al. (2002). Crumbs interacts with moesin and beta(Heavy)-spectrin in the apical membrane skeleton of Drosophila. J. Cell Biol. 158(5): 941-51. 12213838

Neisch, A. L., Formstecher, E. and Fehon, R. G. (2013). Conundrum, an ARHGAP18 orthologue, regulates RhoA and proliferation through interactions with Moesin. Mol Biol Cell 24: 1420-1433. PubMed ID: 23468526

Parlato, S., Giammarioli, A.M., Logozzi, M., Lozupone, F., Matarrese, P., Luciani, F., Falchi, M., Malorni, W. and Fais, S. (2000). CD95 (APO-1/Fas) linkage to the actin cytoskeleton through ezrin in human T lymphocytes: A novel regulatory mechanism of the CD95 apoptotic pathway. EMBO J. 19: 5123-5134. 11013215

Pearson, M. A., Reczek, D., Bretscher, A. and Karplus, P. A. (2000). Structure of the ERM protein moesin reveals the FERM domain fold masked by an extended actin binding tail domain. Cell 101: 259-270. 10847681

Pellikka, M., et al. (1999). Crumbs, the Drosophila homologue of human CRB1/RP12, is essential for photoreceptor morphogenesis. Nature 416(6877): 143-9. 11850625

Pilot, F., Philippe, J. M., Lemmers, C. and Lecuit, T. (2006). Spatial control of actin organization at adherens junctions by a synaptotagmin-like protein Btsz. Nature 442(7102): 580-4. 16862128

Polesello, C., Delon, I., Valenti, P., Ferrer, P. and Payre F. (2002). Dmoesin controls actin-based cell shape and polarity during Drosophila melanogaster oogenesis. Nat. Cell Biol. 4(10): 782-9. 12360288

Roubinet, C., et al. (2011). Molecular networks linked by Moesin drive remodeling of the cell cortex during mitosis. J. Cell Biol. 195(1): 99-112. PubMed Citation: 21969469

Saotome, I., Curto, M. and McClatchey, A. I. (2004). Ezrin is essential for epithelial organization and villus morphogenesis in the developing intestine. Dev. Cell 6(6): 855-64. 15177033

Schottenfeld-Roames, J., Rosa, J. B. and Ghabrial, A. S. (2014). Seamless tube shape is constrained by endocytosis-dependent regulation of active moesin. Curr Biol 24: 1756-1764. PubMed ID: 25065756

Segbert, C., et al. (2004). Molecular and functional analysis of apical junction formation in the gut epithelium of Caenorhabditis elegans. Dev. Biol. 266(1): 17-26. 14729475

Serano, J. and Rubin, G. M. (2003). The Drosophila synaptotagmin-like protein bitesize is required for growth and has mRNA localization sequences within its open reading frame. Proc. Natl Acad. Sci. 100: 13368-13373. 14581614

Shaw, R. J., Henry, M., Solomon, F. and Jacks, T. (1998). RhoA-dependent phosphorylation and relocalization of ERM proteins into apical membrane/actin protrusions in fibroblasts. Mol. Biol. Cell 9: 403-419. 9450964

Sherrard, K. M. and Fehon, R. G. (2015). The transmembrane protein Crumbs displays complex dynamics during follicular morphogenesis and is regulated competitively by Moesin and aPKC. Development 142(10):1869-78. PubMed ID: 25926360

Simons, P. C., Pietromonaco, S. F., Reczek, D., Bretscher, A. and Elias, L. (1998). C-terminal threonine phosphorylation activates ERM proteins to link the cell's cortical lipid bilayer to the cytoskeleton. Biochem. Biophys. Res. Commun. 253: 561-565. 9918767

Speck, O., Hughes, S. C., Noren, N. K., Kulikauskas, R. M. and Fehon, R. G. (2003). Moesin functions antagonistically to the Rho pathway to maintain epithelial integrity. Nature 421(6918): 83-7. 12511959

Takahashi, K., Sasaki, T., Mammoto, A., Takaishi, K., Kameyama, T., Tsukita, S. and Takai, Y. (1997). Direct interaction of the Rho GDP dissociation inhibitor with ezrin/radixin/moesin initiates the activation of the Rho small G protein. J. Biol. Chem. 272: 23371-23375. 9287351

Takahashi, K., et al. (1998). Interaction of radixin with Rho small G protein GDP/GTP exchange protein Dbl. Oncogene 16: 3279-3284. 9681826

Teuliere, J., et al. (2011). MIG-15 and ERM-1 promote growth cone directional migration in parallel to UNC-116 and WVE-1. Development 138(20): 4475-85. PubMed Citation: 21937599

Tran Quang, C., Gautreau, A., Arpin, M. and Treisman, R. (2000). Ezrin function is required for ROCK-mediated fibroblast transformation by the Net and Dbl oncogenes. EMBO J. 19: 4565-4576. 10970850

Tsuda, M., et al. (2004). Crk associates with ERM proteins and promotes cell motility toward hyaluronic acid. J. Biol. Chem. 279(45): 46843-50. 15326184

Ukken, F. P., Aprill, I., JayaNandanan, N. and Leptin, M. (2014). Slik and the receptor tyrosine kinase Breathless mediate localized activation of Moesin in terminal tracheal cells. PLoS One 9: e103323. PubMed ID: 25061859

Van Furden, D., Johnson, K., Segbert, C. and Bossinger, O. (2004). The C. elegans ezrin-radixin-moesin protein ERM-1 is necessary for apical junction remodelling and tubulogenesis in the intestine. Dev. Biol. 272(1): 262-76. 15242805

Verdier, V., et al. (2006). Drosophila Rho-kinase (DRok) is required for tissue morphogenesis in diverse compartments of the egg chamber during oogenesis. Dev. Biol. 297(2): 417-32. Medline abstract: 16887114

Xu, N., Bagumian, G., Galiano, M. and Myat, M. M. (2011). Rho GTPase controls Drosophila salivary gland lumen size through regulation of the actin cytoskeleton and Moesin. Development 138(24): 5415-27. PubMed Citation: 22071107

Yang, Y., et al. (2012). The PP1 phosphatase flapwing regulates the activity of Merlin and Moesin in Drosophila. Dev. Biol. 361(2): 412-26. PubMed Citation: 22133918

Moesin : Biological Overview | Evolutionary Homologs | Regulation | Developmental Biology | Effects of Mutation

date revised: 30 June 2015

Home page: The Interactive Fly © 2003 Thomas B. Brody, Ph.D.

The Interactive Fly resides on the
Society for Developmental Biology's Web server.