α actinin: Biological Overview | References
Gene name - α actinin
Cytological map position - 2C4-2C7
Function - cytoskeleton
Symbol - Actn
FlyBase ID: FBgn0000667
Genetic map position X: 1,919,127..1,936,344 [-]-
Classification - Spectrin repeats, Calponin homology domain
Cellular location - cytoplasmic
α-Actinin is an evolutionarily conserved actin filament crosslinking protein with functions in both muscle and non-muscle cells. In non-muscle cells, interactions between α-actinin and its many binding partners regulate cell adhesion and motility. In Drosophila, one non-muscle and two muscle-specific α-actinin isoforms are produced by alternative splicing of a single gene. In wild-type ovaries, α-actinin is ubiquitously expressed. The non-muscle α-actinin mutant ActnΔ233, which is viable and fertile, lacks α-actinin expression in ovarian germline cells, while somatic follicle cells express α-actinin at late oogenesis. This latter population of α-actinin, termed FC-α-actinin, is shown to be absent from the dorsoanterior follicle cells, and evidence is presented that this is the result of a negative regulation by combined Epidermal growth factor receptor (EGFR) and Decapentaplegic signalling. Furthermore, EGFR signalling increases the F-actin bundling activity of ectopically expressed muscle-specific α-actinin. A novel morphogenetic event in the follicle cells is described that occurs during egg elongation. This event involves a transient repolarisation of the basal actin fibres and the assembly of a posterior β-integrin-dependent adhesion site accumulating α-actinin and Enabled. Clonal analysis using Actn null alleles demonstrated that although α-actinin is not necessary for actin fibre formation or maintenance, the cytoskeletal remodelling is perturbed, and Enabled does not localise in the posterior adhesion site. Nevertheless, epithelial morphogenesis proceeded normally. This work provides the first evidence that α-actinin is involved in the organisation of the cytoskeleton in a non-muscle tissue in Drosophila (Wahlström, 2006).
Throughout development, interactions between the epithelium and the underlying tissue determine the morphology of developing organs. Signals transmitted between the tissues regulate events such as proliferation, apoptosis and cell shape changes. Drosophila oogenesis has long served as a model system for studies concerning patterning and morphogenesis. The structure that supports the development of the oocyte is called an egg chamber. Each egg chamber consists of three cell types: one posteriorly located oocyte, 15 germline-derived nurse cells and a monolayer of somatic follicle cells that envelops the germline cells. The process of oogenesis has been divided into 14 stages, where stage 14 corresponds to a mature egg with a chorion secreted by the follicle cells (Wahlström, 2006).
The shape of the mature egg is dictated by the follicular epithelium, which undergoes several dynamic rearrangements during the later half of oogenesis. During stage 9, a majority of the follicle cells migrate posteriorly over the nurse cells to cover the oocyte. By stage 10, the oocyte has grown to occupy half of the egg chamber, while the nurse cells occupy the other half. During stage 11, the nurse cells transport their cytoplasm into the oocyte in a process called dumping. This results in a rapid increase of the oocyte volume and a concomitant expansion of the follicular epithelium, termed egg elongation (Wahlström, 2006).
Dorsal follicle cell fate is induced at stage 8-9 by Gurken signalling from the oocyte nucleus to the Epidermal growth factor receptor (EGFR) in the overlying follicle cells. This activates an autoregulatory EGFR signalling cascade involving amplification and lateral expansion of the signal and finally downregulation at the dorsal midline. This autoregulatory circuit splits the dorsoanterior domain into two lateral groups of cells that will form the dorsal appendages and one dorsally located group of cells that will contribute to the operculum. The position of the dorsal appendages along the anterior-posterior (A/P) axis of the egg chamber is specified by the TGFβ-family member Decapentaplegic (Dpp), which is secreted by the anterior follicle cells. The two dorsal appendage primordia are composed of roof cells and floor cells. Extensive cell shape changes and a subsequent anterior migration during stages 11-13 result in the formation of two elongated tubes that will secrete the dorsal appendages. In contrast to the wealth of knowledge on patterning molecules involved in all these events, less is known about the cytoskeletal changes directing the morphological rearrangements within the follicular epithelium (Wahlström, 2006).
α-Actinin is an evolutionarily conserved actin crosslinking protein that belongs to the spectrin superfamily (Virel, 2004). In muscle cells, α-actinin is located in the Z-disc, where it crosslinks actin filaments from adjacent sarcomeres (Clark, 2002). In non-muscle cells, α-actinin is localised in stress fibres, lamellipodia, cell–cell and cell–matrix adhesion sites (Otey, 2004). The functional α-actinin molecule is an antiparallel dimer with an actin-binding domain at each end. The monomer is composed of an N-terminal actin-binding domain, four central spectrin-like repeats and a C-terminal calmodulin-like domain with two Ca2+-binding EF-hand motifs (Djinović-Carugo, 1999). In vertebrates, separate genes encode muscle and non-muscle α-actinin isoforms with differing EF-hand sequences. The muscle isoforms lack Ca2+-binding activity, whereas Ca2+ inhibits the actin-binding activity of the non-muscle isoforms (Wahlström, 2006).
Vertebrate α-actinin can directly interact with a large number of different types of molecules in addition to F-actin, suggesting that it plays multiple roles in the cell (Otey, 2004). Most studies on α-actinin’s role in non-muscle cells have focussed on its role in stress fibres and focal adhesions. In the latter, α-actinin interacts directly with several adhesion site components, including β-integrin (Otey, 1990). A mouse knock-out for the major non-muscle α-actinin isoform has not yet been described, but experiments on cultured mammalian cells have suggested that α-actinin is important for stress fibre maintenance (Pavalko, 1991), for maintaining the link between the stress fibre and the adhesion site (Rajfur, 2002) and for adhesion site disassembly (Bhatt, 2002; Wahlström, 2006 and references therein).
In Drosophila, three α-actinin isoforms (non-muscle α-actinin, larval muscle-specific α-actinin and adult muscle-specific α-actinin) are encoded by a single alternatively spliced gene (Fyrberg, 1990; Roulier, 1992). Moreover, the non-muscle isoform is expressed mainly from a promoter different from that of the muscle-specific isoforms (Fyrberg, 1998). The three isoforms differ in the region between the actin-binding domain and the first central repeat. They all share the same C-terminal domain, which according to the sequence was predicted to bind Ca2+ (Fyrberg, 1990; Roulier, 1992). However, no Ca2+-binding activity was detected in vitro (Dubreuil, 1991). α-Actinin null mutants lacking all three isoforms are larval lethal, with defects reported only in muscles (Fyrberg, 1990; Roulier, 1992; Fyrberg, 1998; Wahlström, 2006 and references therein).
In wild-type embryos and ovaries, α-actinin is ubiquitously expressed (Wahlström, 2004). In the viable and fertile mutant ActnΔ233, a small deletion disrupts the first untranslated exon of the non-muscle-specific transcript. The ActnΔ233 mutant lacks detectable expression of α-actinin protein in early embryos, in the ovarian nurse cells and in follicle cells at early stages of oogenesis, yet both embryogenesis and oogenesis proceed normally. In ActnΔ233 mutant ovaries, polar cells and follicle cells from stage 10 onwards still express α-actinin, indicating that α-actinin expression is differently regulated in these cells (Wahlström, 2004). The aim of this study was to examine the population of α-actinin expressed in the ActnΔ233 mutant and to analyse its role in the follicle cells. This study shows that this population of α-actinin, termed FC-α-actinin (follicle cell-α-actinin), is negatively regulated by the EGFR and Dpp signalling pathways. Furthermore, it is shown that α-actinin and β-integrin are required for a previously undescribed cytoskeletal remodelling that occurs in the main body follicle cells during egg elongation (Wahlström, 2006).
To understand how α-actinin is involved in the function of the follicle cells at late oogenesis, α-actinin localisation was studied in wild-type follicle cells at stages 10-14. For detection, used the monoclonal antibody MAC276, which recognises all three α-actinin isoforms (Lakey, 1990; Roulier, 1992) was used, along with a staining protocol that does not allow simultaneous labelling of F-actin with phalloidin. The follicle cells are polarised with the apical side facing the germline and the basal side facing the epithelial sheath surrounding each string of developing egg chambers. At the time of egg chamber assembly, the basal surface of the follicle cells acquires a layer of stress fibre-like actin bundles, which is maintained throughout oogenesis. At stage 10A, α-actinin was localised at the cell cortex (not shown) and was especially abundant in the basal actin fibres. At stage 10B/11, the evenly stained actin fibres began to reorganise, and by stage 11, a distinct patch of α-actinin accumulation was detected in the posterior part of the cell. This pattern was seen in all main body follicle cells, i.e. ventral follicle cells and dorsal cells posterior to the dorsal appendages. In the dorsoanterior follicle cells, α-actinin was expressed at lower levels and showed less distinct localisation patterns. α-Actinin was also detected at the roof cell apices of the elongating dorsal appendages. At the end of stage 12, the basal α-actinin pattern in the main body follicle cells was reorganised again. The accumulation at the posterior end of the cell gradually dispersed, and at stage 13, α-actinin was concentrated at the lateral cell margins. The central actin fibres were less strongly labelled. By stage 14, when the basal actin fibres have disappeared, α-actinin displayed a cortical localisation (Wahlström, 2006).
The lateral stripes of α-actinin in the follicle cells at stage 13 correspond to the previously described adhesion sites shown to contain β-integrin and Ena (Bateman, 2001; Baum, 2001). Integrins are transmembrane receptors for ligands in the extracellular matrix (ECM), and they mediate adhesion between the cell and the ECM (Brown, 2000). Ena is the sole Drosophila member of the conserved family of Ena/VASP proteins, which act as positive regulators of actin filament assembly. Co-localisation studies of α-actinin and Ena revealed a complete overlap in the basal cytoskeleton, including the posterior patch, during stages 11 and 12. At stage 13, there was also extensive co-localisation, although Ena appeared to be located closer to the cell margin than α-actinin. Thus, both α-actinin and Ena accumulate in a transient adhesion site-like structure that forms at the onset of egg elongation (Wahlström, 2006).
The basal stress fibres are aligned perpendicular to the A/P axis of the oocyte between stages 7 and 10, but then a phase of slight disorganisation occurs before the perpendicular alignment is reassumed by stage 13. The disorganised phase correlates with the relocalisation of α-actinin and Ena observed in the basal cytoskeleton. The remodelling could also be recognised by phalloidin-staining of the actin fibres, although they indeed appeared quite irregular in most cells. In several cells, they are polarised in the A/P direction, and they often also converge in a denser patch of F-actin, which overlaps with the posterior patch containing Ena and α-actinin. Thus, egg elongation involves an organised repolarisation of the basal actin fibres (Wahlström, 2006).
Analysis of the α-actinin expression pattern in the non-muscle mutant ActnΔ233 revealed that at least two separate populations of α-actinin are present in the follicle cells (Wahlström, 2004). α-Actinin produced from an mRNA that is transcribed from the upstream promoter (NC-α-actinin) is ubiquitously expressed in the egg chamber. The second α-actinin population, FC-α-actinin, corresponds to α-actinin that is present in certain non-muscle cells of all examined non-muscle-specific α-actinin mutants. FC-α-actinin is most likely produced from an mRNA transcribed from the downstream promoter and may include both non-muscle α-actinin and adult muscle-specific α-actinin. However, an analysis using isoform-specific antibodies or a complete sequencing of the mRNAs expressed in the egg chamber will be required in order to clarify this issue. FC-α-actinin protein was expressed in the main body follicle cells starting from stage 10, but excluded from the dorsoanterior cells. The dorsoanterior cells are patterned by the EGFR and Dpp signalling pathways, and the results showed that these two pathways together downregulate FC-α-actinin expression, but not the expression of NC-α-actinin (Wahlström, 2006).
The dorsoanterior and main body follicle cells undergo very different morphogenetic changes. The dorsoanterior cells elongate in the apicobasal direction and migrate (Dorman, 2004), an event that did not seem to require α-actinin. In contrast, the main body follicle cells flatten and expand their surfaces. These events are expected to involve different sets of cytoskeletal regulators, of which very little is yet known. The formation of a dense layer of basal actin fibres in the main body follicle cells may include upregulation of proteins known to be involved in stress fibre formation, such as α-actinin. It has been shown that the dorsal midline cells upregulate basal E-cadherin and FasIII, indicating increased cell-cell adhesion. These cells also lose their basal actin fibres, which may explain why less α-actinin, i.e., only NC-α-actinin, is expressed in these cells (Wahlström, 2006).
Throughout oogenesis, the basal cytoskeleton is organised into actin fibres aligned in parallel. Variation has been noted in the actin fibre polarity at the late stages of oogenesis. These observations are extended by showing that the basal cytoskeleton undergoes an organised remodelling during the final stages of oogenesis. The rapid increase in oocyte volume during nurse cell dumping at stage 11 requires that the follicle cells expand their surfaces in order to maintain a coherent epithelium. This process involves a transient change in the polarity of the basal actin fibres, from a perpendicular to a parallel orientation relative to the A/P axis of the egg chamber, and the assembly of a transient structure that accumulates α-actinin and Ena. The association of this structure with an accumulation of β-integrin and its dependence on integrin adhesion demonstrate that the cytoskeletal reorganisation is linked to a remodelling of integrin-based adhesion sites. The fact that α-actinin and β-integrin did not show a strict co-localisation is in good agreement with studies on mammalian cells showing that integrin, but not α-actinin, is present in nascent adhesion sites termed focal complexes. α-Actinin accumulation in the adhesion site occurs later, as the focal complexes mature into focal adhesions (Zaidel-Bar, 2003). The signal that induces the remodelling of the basal cytoskeleton remains to be identified. An intriguing possibility is that the mechanical stress applied to the epithelium during nurse cell dumping is transduced into biochemical signals that result in the observed reorganisation. Two different mechanisms are known to mediate mechanotransduction: stretch-activated ion channels or conformational changes within cell-matrix adhesion sites (Wahlström, 2006).
The current view is that the parallel basal actin fibres shape the oocyte during egg elongation by preventing axial expansion. However, since the basal actin fibres repolarise during egg elongation, the current model does not adequately explain how the oocyte acquires its final shape. The fact that Ena accumulates in the posterior of the cell during egg elongation suggests that a mechanism involving localised actin polymerisation and directed cell growth may also contribute to shaping the oocyte. It has been reported that egg elongation is blocked by mutations in the genes encoding α-integrin, β-integrin, the adhesion site components talin or tensin, the receptor tyrosine phosphatase Dlar or the ECM component Laminin A. In the case of β-integrin and Dlar, it has been shown that the actin fibre polarity is disturbed (Bateman, 2001; Frydman, 2001), and this has been suggested to be the cause of the short egg phenotype. However, the data presented in this work give reason to speculate that defective adhesion between the follicle cells and the ECM might play a role as well (Wahlström, 2006).
To explore the function of α-actinin in the main body follicle cells, clones of cells lacking α-actinin were generated. This experiment unexpectedly revealed that α-actinin is not required for the formation or maintenance of the basal actin fibres. Previous studies, relying on the introduction of truncated α-actinin molecules into cultured mammalian cells, have suggested that α-actinin is important for stress fibre maintenance. Furthermore, examination of transformed cells expressing different levels of α-actinin showed that cells with low α-actinin levels had poorly developed stress fibres, an effect was not observe in this study. It is possible that the follicle cell basal actin fibres are not true contractile stress fibres and therefore do not depend on α-actinin. Alternatively, an alternative pathway for stress fibre assembly that is independent of α-actinin might be activated in the follicle cells following removal of α-actinin (Wahlström, 2006 and references therein).
The clonal analysis revealed that while α-actinin was not necessary for the lateral accumulation of Ena at stage 13, it was cell-autonomously required for the posterior localisation of Ena at stages 11 and 12. The reason for this could be that α-actinin is specifically required for recruiting Ena to the posterior adhesion site, perhaps by recruiting their common binding partner zyxin (Drees, 1999). Alternatively, the posterior adhesion site may not form at all. The latter possibility is supported by observations on mosaic stage 10B/11 egg chambers that are in the process of assembling the posterior adhesion site. While wild-type cells are in the process of translocating Ena towards the posterior, neighbouring cells lacking α-actinin still showed a lateral Ena pattern. At stage 12/13, the lateral adhesion sites are assembled earlier in the mutant cells than in the wild-type cells, perhaps because the mutant cells had not reorganised their cytoskeleton to the same extent as the wild-type cells had. Thus, these results clearly demonstrate that adhesion site remodelling is altered in the absence of α-actinin. However, in contrast to the cells lacking β-integrin, the Actn mutant cells appear to maintain their adhesion to the ECM, since they appear equally well spread as the wild-type cells at stage 13 (Wahlström, 2006).
The results are in agreement with the current view that vertebrate α-actinin is involved in adhesion site disassembly. This is a strictly regulated process that involves signalling by phosphoinositides, tyrosine phosphorylation and proteolytic cleavage of individual components (Carragher, 2004). α-Actinin is one of the targets for these activities. Phosphorylation of α-actinin by focal adhesion kinase (FAK) reduces α-actinin’s affinity for F-actin and regulates the activity of FAK itself (Izaguirre, 2001; Zhang, 2006), PtdIns(3,4,5)-P3 binding to α-actinin disrupts α-actinin binding to β-integrin and F-actin (Greenwood, 2000; Fraley, 2005), and cleavage of α-actinin by calpain has been associated with cell shape changes in certain cell types. It has also been shown that α-actinin is essential for maintaining the link between the adhesion site and the stress fibre. This conclusion was reached based on an experiment showing that laser-mediated inactivation of α-actinin located in an adhesion site resulted in stress fibre detachment from the adhesion site (Rajfur, 2002). In the Drosophila follicle cells, α-actinin is clearly not required for actin fibre attachment. However, by the laser-mediated inactivation, α-actinin was removed from an adhesion site, whereas in the Actn null mutant follicle cells, the adhesion sites never contained α-actinin. Considering the large number of proteins that interact with α-actinin, it is expected that a signal targeted at α-actinin indirectly affects many other proteins and processes as well. An adhesion site lacking α-actinin may well be functional, but it may respond differently to various signals that induce adhesion site remodelling (Wahlström, 2006).
Interestingly, even very large clones of Actn mutant cells had no negative effects on egg morphology. This indicates that proper cytoskeletal remodelling and posterior localisation of Ena is not necessary for egg elongation. Apparently, the expansion of the main body follicle cells in the Actn mutant cells occurs by an alternative mechanism that is not dependent on α-actinin. This raises the question of whether the wild-type remodelling mechanism would become important under some specific conditions not prevailing in the laboratory. The impact of the environment on the development of mutant phenotypes has been well documented in the slime mould Dictyostelium discoideum. Lack of α-actinin results in only minor alterations in cellular functions and did not reduce viability. However, when the cells were grown under conditions resembling their natural habitat, specific developmental defects appeared (Wahlström, 2006).
The dorsoanterior follicle cells express only NC-α-actinin, whereas the main body follicle cells also express FC-α-actinin. Furthermore, these two cell populations responded differently to overexpression of the adult muscle-specific α-actinin isoform. Prominent actin spikes were induced in the dorsal appendage cells, while the main body follicle cells remained unaffected. This effect might be specific for the adult muscle-specific isoform. Alternatively, the dorsal appendage cells might be sensitive to an overload of any isoform of α-actinin. In either case, it provides a plausible explanation as to why FC-α-actinin needs to be downregulated in these cells. The appearance of spikes roughly coincided with the start of tube elongation, suggesting a connection with events that regulate dorsal appendage migration. At stages 9 and 10A, also stages at which the follicle cells undergo migration, a thickening of the basal actin fibres was observed. Since the dorsal appendage cells are characterised by their maintained EGFR signalling, whether EGFR signalling directly increased α-actinin's bundling activity was tested. Indeed, co-expression of λtop and muscle-specific α-actinin at stages 9 and 10A resulted in extensive bundling of the basal actin fibres, an effect that was not observed when either protein was expressed alone. No effect was seen prior to stage 9, suggesting that the factor needed for α-actinin’s bundling activity was induced independent of EGFR signalling at stage 9, but subsequent to the induction, its activity was modulated by EGFR signalling (Wahlström, 2006).
Regardless of whether or not the effect is specific for a certain α-actinin isoform, it is concluded that EGFR signalling modulates α-actinin's bundling activity. A possible mediator could be, for example, an enzyme that regulates the phosphoinositide levels in the cell. The F-actin crosslinking activity of vertebrate α-actinin is regulated by phosphoinositide binding, although some controversy exists as to whether phosphoinositides increase (Fukami, 1992) or decrease (Fraley, 2003) the crosslinking activity. Interestingly, mammalian cells transfected with a mutant form of α-actinin-1 with a reduced affinity for phosphoinositides displayed excessive bundling of actin filaments (Fraley, 2003) in a manner somewhat similar to what was observed in stage 9 follicle cells overexpressing both λtop and muscle-specific α-actinin. The phosphoinositide binding site in vertebrate α-actinin has been mapped to the N-terminal actin-binding domain. All residues shown to be important for phosphoinositide binding (Fukami, 1996; Fraley, 2003; Franzot, 2005) are conserved in Drosophila α-actinin (Fyrberg, 1990), indicating that the same regulatory mechanism probably exists in Drosophila as well (Wahlström, 2006).
This study contributes new data to the field of cytoskeletal dynamics in Drosophila follicle cells. A surprisingly complex regulation was undercovered underlaying α-actinin expression in the follicle cells. The basal cytoskeleton of the main body follicle cells undergoes an organised remodelling during egg elongation, and α-actinin is required in this process. This observation provides the first identified phenotype in a Drosophila non-muscle tissue lacking α-actinin. The fact that both loss of α-actinin and overexpression of α-actinin results in very distinct cellular phenotypes suggests that the follicular epithelium could serve as a very useful in vivo system for further studies on mechanisms that regulate α-actinin function and activity. Furthermore, the cytoskeletal remodelling may provide an easily accessible and genetically tractable model for studies on adhesion dynamics in vivo (Wahlström, 2006).
The single copy Drosophila alpha-actinin gene is alternatively spliced to generate three different isoforms that are expressed in larval muscle, adult muscle and non-muscle cells, respectively. Novel alpha-actinin alleles, which specifically remove the non-muscle isoform, have been generated. Homozygous mutant flies are viable and fertile with no obvious defects. Using a monoclonal antibody that recognizes all three splice variants, alpha-actinin distribution was compared in wild type and mutant embryos and ovaries. Non-muscle alpha-actinin is present in young embryos and in the embryonic central nervous system. In ovaries, non-muscle alpha-actinin is localized in the nurse cell subcortical cytoskeleton, cytoplasmic actin cables and ring canals. In the mutant, alpha-actinin expression remains in muscle tissues, but also in a subpopulation of epithelial cells in both embryos and ovaries. This suggests that various populations of non-muscle cells regulate alpha-actinin expression in different ways. Ectopically expressed adult muscle-specific alpha-actinin localizes to all F-actin containing structures in the nurse cells in the absence of endogenous non-muscle alpha-actinin (Wahlström, 2006; full text of article).
The rolling pebbles gene of Drosophila encodes two proteins, one of which, Rols7, is essential for myoblast fusion. In addition, Rols 7 is expressed during myofibrillogenesis and in the mature muscles. Here it overlaps with alpha-Actinin (a-Actn) and the N-terminus of D-Titin/Kettin/Zormin in the Z-line of the sarcomeres. In the attachment sites of the somatic muscles, Rols7 and the immunoglobulin superfamily protein Dumbfounded/Kin of irreC (Duf/Kirre) colocalise. As Duf/Kirre is detectable only transiently, it may be involved in establishing the first contact of the outgrowing muscle fiber to the epidermal attachment site. It is proposed that Rols7 and Duf/Kirre link the terminal Z-disc to the cell membrane by direct interaction. This is supported by the fact that in yeast two hybrid assays the tetratricopeptide repeat E (TPR E) of Rols7 shows interaction with the intracellular domain of Duf/Kirre. The colocalisation of Rols7 with a-Actn and with D-Titin/Kettin/Zormin in the Z-dics is reflected in interactions with different domains of Rols7 in this assay. In summary, these data show that besides the role in myoblast fusion, Rols7 is a scaffold protein during myofibrillogenesis and in the Z-line of the sarcomere as well as in the terminal Z-disc linking the muscle to the epidermal attachment sites (Kreiskother, 2006).
The scaffold protein Rols7 has been shown to be essential for myoblast fusion in the somatic mesoderm during Drosophila embryogenesis where it might interact with several components of the fusion machinery. Evidence is presented that Rols7 has an additional function in the establishment of the muscle attachment and the formation of the Z-discs, as well as in the Z-discs of the mature muscles (Kreiskother, 2006).
During myoblast fusion, Rols7 mRNA decays at stage 15. Antibody staining of stage 17 embryos reveal a concentration of Rols7 at the muscle ends next to the epidermal attachment sites, which are caused by new transcription in RT-PCR experiments. Later on in the mature larval muscles Rols7 is detected in the sarcomeric Z-discs (Kreiskother, 2006).
During the early stages of myogenesis, the interaction of the founder cell specific protein Duf/Kirre and the fusion competent myoblasts (fcm) specific Sns leads to the adhesion of the two cell types, which is a prerequisite for further steps of the fusion process. Besides this, Duf/Kirre transiently are concentrated at the end of the developing muscles at stage 15 and 16, while it disappears again at stage 17. This led to a hypothesis that Duf/Kirre might participate in the first contact of the outgrowing muscle to the attachment site, as does Vein. This would require an interaction partner in the extracellular matrix or at the epidermal site. Since the sns transcript is present in the muscle attachment sites at a low level at stage 17, antibody staining for Sns was performed, but a distinct signal in the attachment sites could not be detected. As well as the transcript of sns, its paralog, Hibris (Hbs), is also found in the muscle attachment sites, and, more exactly, localised to the contact site between the cells at the epidermal attachments. Thus, it could function as an interaction partner for Duf/Kirre (Kreiskother, 2006).
As a further possible interaction partner Rst/IrreC was considered, since Rst/IrreC, the paralogue of Duf/Kirre, shows expression in the epidermal tendon cells during embryonic stages. Due to the fact that Duf/Kirre and Rst/IrreC are indeed able to undergo heterophilic interaction in cell culture experiments, the conclusion is drawn that Rst/IrreC might be the candidate for an interaction partner of Duf/Kirre on the epidermal site, thus enabling an early contact of the muscle to the epidermal attachment site (Kreiskother, 2006).
Rols7, which interacts with the intracellular domain of Duf/Kirre, is also localised at the muscle ends from late stage 16 onwards shortly before Duf/Kirre disappears. It is speculated that Rols7 is brought to the membrane where it interacts with Duf/Kirre (Kreiskother, 2006).
alpha actinin (α-Actn) and D-Titin/Kettin, both found at the muscle attachment site in a similar pattern, also interact with Rols7 (at least in the yeast two hybrid assay) and participate in the establishment of the terminal Z-disc. For the flight muscle it was shown that α-Actn is essential for the formation of this structure and for obtaining a correct insertion of the myofibril to the epidermal tendon cell. Furthermore the yeast assay showed an interaction of α-Actn with Duf/Kirreintra (Kreiskother, 2006).
kettin mutants show strong defects in terminal Z-disc function. This study proposes that, in addition to Kettin, Rols7 and α-Actn are important for the formation of this structure. The process might be connected to Muscleblind (Mbl), since in mutants for mbl, Z-discs are not assembled correctly. Unfortunately, a mutant analysis of Rols7 function in terminal Z-disc formation is difficult due to its essential function during myoblast fusion (Kreiskother, 2006).
Apart from the myoblasts and attachment sites, Rols7 is expressed in the developing sarcomeres of larval and adult muscles and localises to the Z-discs, as was shown using antibodies for α-Actn and D-Titin/Kettin as markers. Yeast interaction assays revealed that Rols7 might directly interact with α-Actn and Zormin, which, like Kettin, is an isoform derived from the sallimus (sls) gene and also localises to the Z-discs. Therefore, it is postulated that Rols7 serves as a scaffold protein that links α-Actn and Zormin in the Z-disc. Furthermore, the analyses of alpha actinin mutants showed that the presence of α-Actn is not necessary for Rols7 localisation to the Z-discs. In addition, Rols7, as well as α-Actn and D-Titin/Kettin, is present during the assembly of the sarcomere. In vertebrates it has been shown that in spreading edges of rat cardiomyocytes, dense bodies that contain Z-disc proteins assemble at the spreading membrane and align to premyofibrils in cooperation with newly formed actin filaments and small myosin filaments (Kreiskother, 2006).
Antibody staining showed protein aggregates that aligned to form kinds of premyofibrils and demonstrated that in Drosophila, the assembly of the Z-discs seems to be similar to that of vertebrates. So, Rols7 is the first protein that is essential for myoblast fusion and plays an additional role in the sarcomere assembly as well as in the Z-discs of mature muscles, where it is proposed that it links α-Actn and D-Titin/Kettin/Zormin. Δ-titin/kettin-mutants have a weaker fusion phenotype than rols7-mutants, however, D-Titin/Kettin is clearly expressed during myoblast fusion as a component of the adhesion complex between founder cell and fcm. Individual Rols7 domains serve different function in distinct processes of myogenesis (Kreiskother, 2006).
From coimmunoprecipitation experiments it was already supposed that the intracellular domain of Duf interacts with Rols. Furthermore, cell culture cotransfection assays showed colocalisation of Duf, Rols and D-Titin. In yeast interaction assays the individual domains of Rols7 were tested for interaction with potential partners that included components of the fusion machinery which might be relevant for muscle attachment or sarcomere assembly as well. Indeed, the different domains interact with different partners in different developmental contexts, and it is concluded that Rols7 is a multifunctional protein (Kreiskother, 2006).
The interaction of Rols7 with the intracellular domain of Duf/Kirre was confirmed and it was found that the interaction probably is mediated by the TPR repeats of Rols7, respectively, by the most C-terminal TPR E repeat and the R1 fragment that contains the RING finger and an additional part of 321 amino acids downstream. In contrast α-Actn interacts with the R1 domain and with both TPR repeats, the TPR E and the TPR X, whereas the N-terminal part of Zormin interacts only with the R1 domain in this assay. No interaction was detected for the N-terminal part of Kettin and the Rols7 domains. These results, together with the rescue capability of truncated Rols7 versions, led to the proposal of certain functions to individual Rols7 domains. Either the RING finger domain, the TPR repeats or the ankyrin repeats and the TPR repeats have been deleated and the remaining parts of Rols7 were examined for their competence to rescue the rols fusion defect. A deletion of the RING finger domain does not affect the rescue of the rols fusion phenotype, whereas a deletion of the TPR repeats leads to a partial rescue and a deletion of ankyrin repeats and TPR repeats together does not rescue fusion at all. The Rols7 version without the RING finger rescues the fusion phenotype. This RING finger is included in the R1 fragment which interacts with Duf/Kirreintra, Blow, Zormin and α-Actn. Thus, it is proposed that the R1 domain is a candidate to mediate the transient interaction of Duf/Kirreinttra at the muscle attachment sites. The R1 domain of Rols7 could then mediate the interaction with Zormin in the Z-discs in all larval muscles. R1 is the only Rols7 fragment that interacts with Zormin. R1 and TPR E as well as TPR X have the capability to interact with α-Actn. It cannot be decide whether both domains of Rols7 interact with α-Actn in the Z-discs. The interaction of Duf/Kirreintra with the TPR E repeat indicates a function of the TPR E repeat during myoblast fusion, since its deletion only leads to a partial rescue of the rols fusion phenotype. The ankyrin repeats did not interact with any of the proteins which have been tested in the yeast assay and which are characteristic for sarcomere assembly and muscle attachment. Taking this together with the fact that a deletion of this domain, in addition to a deletion of the TPR repeats, prevents the rescue of the fusion defect, indicates that the ankyrin repeats predominantly function during myoblast fusion (Kreiskother, 2006).
Rols7 is a scaffold protein which contains distinct domains characteristic of protein-protein interaction. It is proposed that the interaction of the appropriate domain with certain proteins is specific for the process of myogenesis, myoblast fusion, muscle attachment or sarcomere assembly (Kreiskother, 2006).
Drosophila Muscleblind (Mbl) proteins control terminal muscle and neural differentiation, but their molecular function has not been experimentally addressed. Such an analysis is relevant as the human Muscleblind-like homologs (MBNL1-3) are implicated in the pathogenesis of the inherited muscular developmental and degenerative disease myotonic dystrophy. The Drosophila muscleblind gene expresses four protein coding splice forms (mblA to mblD) that are differentially expressed during the Drosophila life cycle, and which vary markedly in their ability to rescue the embryonic lethal phenotype of muscleblind mutant flies. Analysis of muscleblind mutant embryos reveals misregulated alternative splicing of the transcripts encoding Z-band component alpha-Actinin, which can be replicated in human cells expressing a Drosophila alpha-actinin minigene and epitope-tagged Muscleblind isoforms. MblC appreciably altered alpha-actinin splicing in this assay, whereas other isoforms had only a marginal or no effect, demonstrating functional specialization among Muscleblind proteins. To further analyze the molecular basis of these differences, the subcellular localization of Muscleblind isoforms was studied. Consistent with the splicing assay results, MblB and MblC were enriched in the nucleus while MblA was predominantly cytoplasmic. In myotonic dystrophy, transcripts bearing expanded non-coding CUG or CCUG repeats interfere with the function of human MBNL proteins. Co-expression of CUG repeat RNA with the alpha-actinin minigene altered splicing compared with that seen in muscleblind mutant embryos, indicating that CUG repeat expansion RNA also interferes with Drosophila muscleblind function. Moreover MblA, B, and C co-localize with CUG repeat RNA in nuclear foci in cell culture. These observations indicate that Muscleblind isoforms perform different functions in vivo, that MblC controls muscleblind-dependent alternative splicing events, and establish the functional conservation between Muscleblind and MBNL proteins both over a physiological target (alpha-actinin) and a pathogenic one (CUG repeats) (Vicente, 2007).
Search PubMed for articles about Drosophila alpha-actinin
Bateman, J., Reddy, R. S., Saito, H. and Van Vactor, D. (2001). The receptor tyrosine phosphatase Dlar and integrins organize actin filaments in the Drosophila follicular epithelium. Curr. Biol. 11: 1317-1327. PubMed citation: 11553324
Baum, B. and Perrimon, N. (2001). Spatial control of the actin cytoskeleton in Drosophila epithelial cells. Nat. Cell Biol. 3: 883-890. PubMed citation: 11584269
Bhatt, A., Kaverina, I., Otey, C. and Huttenlocher, A. (2002). Regulation of focal complex composition and disassembly by the calcium-dependent protease calpain. J. Cell Sci. 115: 3415-3425. PubMed citation: 12154072
Brown, N. H., Gregory, S. L. and Martin-Bermudo, M.D. (2000). Integrins as mediators of morphogenesis in Drosophila. Dev. Biol. 223: 1-16. PubMed citation: 10864456
Carragher, N.O. and Frame, M.C. (2004). Focal adhesion and actin dynamics: a place where kinases and proteases meet to promote invasion. Trends Cell Biol. 14: 241-249. PubMed citation: 15130580
Clark, K. A., et al. (2002). Striated muscle cytoarchitecture: an intricate web of form and function. Annu. Rev. Cell Dev. Biol. 18: 637-706. PubMed citation: 12142273
Djinović-Carugo, K., Young, P., Gautel, M. and Saraste, M. (1999). Structure of the α-actinin rod: molecular basis for cross-linking of actin filaments. Cell 98: 537-546. PubMed citation: 10481917
Dorman, J. B. et al. (2004). bullwinkle is required for epithelial morphogenesis during Drosophila oogenesis, Dev. Biol. 267: 320-341. PubMed citation: 15013797
Drees, B. E., Andrews, K. M. and Beckerle, M. C. (1999). Molecular dissection of zyxin function reveals its involvement in cell motility. J. Cell Biol. 147: 1549-1560. PubMed citation: 10613911
Dubreuil, R. R. et al. (1991). Structure, calmodulin-binding, and calcium-binding properties of recombinant αspectrin polypeptides. J. Biol. Chem. 266: 7189-7193. PubMed citation: 2016322
Fraley, T. S., et al. (2003). Phosphoinositide binding inhibits α-actinin bundling activity. J. Biol. Chem. 278: 24039-24045. PubMed citation: 12716899
Fraley, T. S., et al. (2005). Phosphoinositide binding regulates α-actinin dynamics: mechanism for modulating cytoskeletal remodeling. J. Biol. Chem. 280: 15479-15482. PubMed citation: 15710624
Franzot, B., Sjöblom, B., Gautel, M. and Djinovic Carugo, K. (2005). The crystal structure of the actin binding domain from α-actinin in its closed conformation: structural insight into phospholipid regulation of α-actinin. J. Mol. Biol. 348: 151-165. PubMed citation: 15808860
Frydman, H. M. and Spradling, A. C. (2001). The receptor-like tyrosine phosphatase Lar is required for epithelial planar polarity and for axis determination within Drosophila ovarian follicles. Development 128: 3209-3220. PubMed citation: 11688569
Fukami, K., et al. (1992). Requirement of phosphatidylinositol 4,5-bisphosphate for α-actinin function. Nature 359: 150-152. PubMed citation: 1326084
Fukami, J., et al. (1996). Identification of a phosphatidylinositol 4,5-bisphosphate-binding site in chicken skeletal muscle α-actinin, J. Biol. Chem. 271: 2646-2650. PubMed citation: 8576235
Fyrberg, E., et al. (1990). Molecular genetics of Drosophila alpha-actinin: mutant alleles disrupt Z disc integrity and muscle insertions. J. Cell Biol. 110: 1999-2011. PubMed citation: 2112549
Fyrberg, C., et al. (1998).Characterization of lethal Drosophila melanogaster α-actinin mutants, Biochem. Genet. 36: 299-310. PubMed citation: 9919356
Greenwood, J. A., et al. (2000). Restructuring of focal adhesion plaques by PI 3-kinase: regulation by PtdIns (3,4,5)-P3 binding to α-actinin. J. Cell Biol. 150: 627-642. PubMed citation: 10931873
Izaguirre, G., et al. (2001). The cytoskeletal/non-muscle isoform of α-actinin is phosphorylated on its actin-binding domain by the focal adhesion kinase. J. Biol. Chem. 276: 28676-28685. PubMed citation: 11369769
Kreiskother, N., Reichert, N., Buttgereit, D., Hertenstein, A., Fischbach, K. F. and Renkawitz-Pohl, R. (2006). Drosophila rolling pebbles colocalises and putatively interacts with alpha-Actinin and the Sls isoform Zormin in the Z-discs of the sarcomere and with Dumbfounded/Kirre, alpha-Actinin and Zormin in the terminal Z-discs. J. Muscle Res. Cell Motil. 27(1): 93-106. PubMed citation: 16699917
Lakey, A., et al. (1990). Identification and localization of high molecular weight proteins in insect flight and leg muscle. EMBO J. 9: 3459-3467. PubMed citation: 2209553
Otey, C. A., Pavalko, F. M. and Burridge, K. (1990). An interaction between α-actinin and the β1 integrin subunit in vitro. J. Cell Biol. 111: 721-729. PubMed citation: 2116421
Otey, C. A. and Carpen, O. (2004). Alpha-actinin revisited: a fresh look at an old player. Cell Motil. Cytoskeleton 58(2): 104-11. PubMed ID: 15083532
Pavalko, F. M. and Burridge, K. (1991). Disruption of the actin cytoskeleton after microinjection of proteolytic fragments of α-actinin. J. Cell Biol. 114: 481-491. PubMed citation: 1907287
Rajfur, Z., et al. (2002). Dissecting the link between stress fibres and focal adhesions by CALI with EGFP fusion proteins. Nat. Cell Biol. 4: 286-293. PubMed citation: 11912490
Roulier, E. M., Fyrberg, C. and Fyrberg, E. (1992). Perturbations of Drosophila α-actinin cause muscle paralysis, weakness, and atrophy but do not confer obvious nonmuscle phenotypes, J. Cell Biol. 116: 911-922. PubMed citation: 1734023
Vicente, M., et al. (2007). Muscleblind isoforms are functionally distinct and regulate alpha-actinin splicing. Differentiation 75(5): 427-40. PubMed citation: 17309604
Virel, A. and Backman, L. (2004). Molecular evolution and structure of alpha-actinin. Mol. Biol. Evol. 21(6): 1024-31. PubMed ID: 15014165
Wahlström, G., et al. (2004). Drosophila non-muscle α-actinin is localized in nurse cell actin bundles and ring canals, but is not required for fertility. Mech. Dev. 121: 1377-1391. PubMed citation: 15454267
Wahlström, G., Norokorpi, H. L. and Heino, T. I. (2006). Drosophila alpha-actinin in ovarian follicle cells is regulated by EGFR and Dpp signalling and required for cytoskeletal remodelling. Mech. Dev. 123(11): 801-18. PubMed citation: 17008069
Zaidel-Bar, R., Ballestrem, C. Kam, Z. and Geiger, B. (2003). Early molecular events in the assembly of matrix adhesions at the leading edge of migrating cells. J. Cell Sci. 116: 4605-4613. PubMed citation: 14576354
Zhang, Z., Lin, S. Y., Neel, B. G. and Haimovich, B. (2006). Phosphorylated α-actinin and protein-tyrosine phosphatase 1B coregulate the disassembly of the focal adhesion kinase × Src complex and promote cell migration. J. Biol. Chem. 218: 1746-1754. PubMed citation: 16291744
date revised: 15 May 2008
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