The assembly and maintenance of the muscle sarcomere requires a complex interplay of actin- and myosin-associated proteins. Myotilin is a thin filament-associated Z-disc protein that consists of two Ig-domains flanked by a unique serine-rich amino-terminus and a short carboxy-terminal tail. It binds to α-actinin and filamin c and is mutated in limb girdle muscular dystrophy 1A (LGMD1A). Here we show that myotilin also directly binds F-actin, efficiently cross-links actin filaments alone or in concert with α-actinin and prevents filament disassembly induced by Latrunculin A. Myotilin forms dimers via its carboxy-terminal half, which may be necessary for the actin-bundling activity. Overexpression of full-length myotilin but not the carboxy-terminal half induces formation of thick actin cables in non-muscle cells devoid of endogenous myotilin. The expression of myotilin in muscle cells is tightly regulated to the later stages of in vitro myofibrillogenesis, when preassembled myofibrils begin to align. Expression of either amino- or carboxy-terminally truncated myotilin fragments but not wild-type myotilin in differentiating myocytes leads to myofibril disarray. The disease association and functional characteristics indicate an indispensable role for myotilin in stabilization and anchorage of thin filaments, which may be a prerequisite for correct Z-disc organization.
Actin is one of the most abundant proteins in eukaryotic cells and plays a key role in many cellular processes, including cell movement, cytokinesis and muscle cell contraction. It exists as a monomer that readily polymerizes into long, polar filaments. A great number of actin-binding proteins regulate filament stability and length (1,2). Proteins may bind to actin filaments by using several distinct domains like, for instance, calponin-homology (CH) domains found in number of different protein families (3,4) or C2 type immunoglobulin- (Ig-) like domains in proteins like titin and kettin (5,6). The ability to cross-link actin filaments requires at least two actin-binding sites in a single protein (e.g. fascin, fimbrin and espin) (3,7,8). Alternatively, cross-linking can occur via a single actin-binding site if the protein is able to dimerize (9,10). The latter family of actin cross-linkers may be divided further into two distinct subclasses that are defined by the structural motifs responsible for protein dimerization.
In cross-striated muscles, actin filaments form the basis of thin myofilaments, which appear to be relatively static and strikingly precise in their length. They are covered by tropomyosin throughout their length (9,10), capped by tropomodulin at the pointed ends (11) and by capZ at the barbed ends (12). Actin filaments have been proposed to be cross-linked at the Z-disc by Z-filaments, which are thought to consist primarily of α-actinin dimers (13). Nebulin, a large actin-binding protein spanning the entire length of a thin filament, might act as a ruler regulating sarcomeric thin filament length (14–16). On the other hand, the giant muscle protein titin works both as a spring and as a ruler for sarcomere formation (17–19). In addition to α-actinin, titin and nebulin, several other recently characterized structural proteins, including telethonin (also called T-Cap) (20,21), filamin c (formerly called γ-filamin, ABPL, FLN2) (22), FATZ (23) (also called myozenin or calsarcin) (24,25), ZASP (26) and myotilin (27), are located in the Z-disc. These proteins form multiple interactions (e.g. α-actinin binds to titin, FATZ, ZASP and myotilin, filamin c binds to FATZ and myotilin, FATZ binds to telethonin and ZASP), thereby creating a complex protein network, which apparently is required for structural stability and in order to withstand the force generated by muscle contraction. It is still largely unclear how the individual molecules are involved in the formation, maintenance and regulation of the Z-disc.
Myocyte differentiation involves a series of events, during which (i) mononuclear myoblasts withdraw from the cell cycle, (ii) cells fuse into multinuclear myotubes and (iii) myofibrils form, which finally leads to functional, cross-striated myocytes (28–30). During early myofibrillogenesis in vitro, titin and α-actinin appear in a punctuate pattern along stress fibre-like structures (28,29,31). While the amino-terminal portion of titin is already tethered to Z-disc primordia, the remainder of the molecule gets stretched out only subsequently. Once the carboxy-termini of titin molecules from opposing Z-discs meet in the centre, i.e. in the future M-band, they are linked by myomesin (32). This structure seems to form a template for thick filament integration into the sarcomeres (31,33,34). Other thin filament proteins like tropomodulin, nebulin, filamin c and telethonin (22,35,36) are also expressed at early stages of differentiation and they are incorporated into forming premyofibrils, which are later aligned to form mature myofibrils (37).
We recently discovered myotilin, a thin-filament associated protein, which binds to both α-actinin and filamin c (27,38). It is composed of a unique, serine-rich amino-terminus and a carboxy-terminus that contains two Ig-like domains, most homologous to Ig-domains of palladin, myopalladin and Z-disc Ig-domains 7 and 8 of titin (27,39–41). The α-actinin binding site resides in the amino-terminus, whereas sites for filamin c binding and for putative self-association locate at the carboxy-terminus (27,38,42). In adult tissues, myotilin is mainly expressed in skeletal and cardiac muscles and in the peripheral nerves (27). Myotilin missense mutations have been found to cause limb girdle muscular dystrophy 1A (LGMD1A), a dominantly inherited disease (42,43). Clinical features of this disease are weakness of the proximal muscles, elevated creatine kinase levels and reduced deep tendon reflexes in early adulthood (44). Morphological alterations include excessive streaming of Z-lines and presence of filamentous aggregates resembling nemaline rods, degeneration of myofibres, fibre splitting and rimmed vacuoles (42).
The disease association, the myofibrillar abnormalities in LGMD1A patients and the interactions of myotilin with structural components of the sarcomere suggest an important role for myotilin in the organization and/or maintenance of sarcomeres. In this study, we have analyzed the expression and location of myotilin in mature and differentiating myocytes, the impact of myotilin on F-actin organization and the role of myotilin during sarcomere assembly. The results indicate that myotilin is a strong actin cross-linking protein of the Z-disc acting in concert with α-actinin. Its properly timed expression at a late stage of myofibrillogenesis is important for correct assembly of the contractile apparatus.
Myotilin cross-links actin-filaments
We previously reported myotilin to be an α-actinin binding protein, located in the Z-discs of striated muscles (27), where thin filaments of neighboring sarcomeres are anchored. To investigate whether myotilin might have a more direct role in the anchorage of thin filaments, we investigated the capability of myotilin to bind directly to F-actin. Several recombinant fragments, containing portions of either α-actinin or myotilin (see Fig. 1A), were used in combination with actin for in vitro binding assays. In control experiments, F-actin was found to sediment by high-speed centrifugation, while α-actinin and the GST-tagged myotilin remained mostly in solution (Fig. 1B).
Actin filament cosedimentation assays demonstrated that both full-length myotilin (data not shown) and the construct Myot 80–462 (lacking the amino-terminal 79 and the carboxy-terminal 25 amino acids) cosedimented with actin filaments with similar efficiency to α-actinin (Fig. 1B). These experiments demonstrate that myotilin binds actin filaments; however, it does not affect the critical concentration of actin polymerization (data not shown). Cosedimentation assays with different ratios of myotilin and actin filaments showed that maximal binding was reached with a 1:1 stoichiometry of myotilin and actin (Fig. 1C). Interestingly, coincubation of myotilin with actin and α-actinin increased the amount of sedimented α-actinin in comparison to experiments where α-actinin alone was incubated and centrifuged with actin filaments, suggesting that myotilin and α-actinin interact with each other under these conditions (Fig. 1B).
The ability of myotilin to cross-link actin filaments was subsequently tested by low-speed centrifugation. Under these conditions F-actin alone is retained in the supernatant (Fig. 1D), and only cross-linked actin bundles are precipitated. α-actinin, known to cross-link actin filaments, precipitated in this assay most of the F-actin, whereas repeats R1–R4 (lacking the actin-binding site) had no effect on actin-crosslinking and therefore failed to sediment actin (Fig. 1D). Myotilin precipitated actin with an efficiency similar to that of α-actinin, indicating that myotilin is capable of cross-linking actin filaments (Fig. 1D and E). Again, coincubation of myotilin with actin—α-actinin bundles significantly enhanced the amount of α-actinin precipitated (Fig. 1D).
Previously performed yeast two-hybrid experiments had suggested that myotilin is able to dimerize (27). To verify this finding, we expressed a GST-tagged C-terminal fragment (Myot 229–442), cleaved from GST with thrombin and purified using cation exchange chromatography. The purified protein was analyzed on a Superdex 10/30 HR column, where two peaks of ∼25 and ∼50 kDa were separated (Fig. 2A), presumably representing monomeric and dimeric forms of the protein, respectively. Samples from both peaks were subjected to SDS–PAGE and western blotting, and detected by silver staining and immunoblotting using myotilin antibody (Fig. 2B). Interestingly, both peaks contained the 25 kDa myotilin fragment. The identity of the polypeptides in both peaks was further verified by mass spectroscopy (Fig. 2C) and amino-terminal sequencing. These results indicate that myotilin can dimerize in vitro via its carboxy-terminal Ig-domain containing region. The actin filaments and bundles obtained after coincubation with α-actinin and/or myotilin, were also studied by transmission electron microscopy. Actin alone formed thin, long filaments, loosely distributed throughout the grids (Fig. 3A). When α-actinin was mixed with F-actin, the filaments were packed into thick, stiff bundles (Fig. 3B). In contrast, bundles cross-linked by myotilin appeared somewhat looser and more curved (Fig. 3C). A simultaneous incubation of F-actin with both cross-linkers resulted in larger, more tightly packed actin bundles (Fig. 3D), suggesting the assembly of a ternary complex between these three proteins.
Expression of myotilin in eukaryotic cells leads to reorganization of the actin cytoskeleton
To study the in vivo function of myotilin, we transiently expressed full-length myotilin (wt), a truncated HA-tagged myotilin fragment comprising amino acids 215–498, or β-galactosidase (as a control) in COS-1 and CHO cells. Transfection of the control cDNA in both cell types gave a diffuse cytoplasmic β-galactosidase staining (Fig. 4). The carboxy-terminus of myotilin colocalized with the cortical actin cytoskeleton as visualized by phalloidin co-staining (Fig. 4). This suggests an association between myotilin and F-actin also in vivo. Overexpression of this part of the protein, however, had no effect on F-actin organization. Full-length myotilin also colocalized with F-actin in both cell types. Interestingly, in COS-1 cells, the overexpressed polypeptide induced formation of prominent actin cables and caused a dramatic reorganization of the actin cytoskeleton. Similarly, the transfection of CHO-cells, which have a more organized actin cytoskeleton in comparison to COS-1 cells, induced the formation of prominent actin cables. In CHO cells, the recombinant protein appeared in a periodic punctate pattern along actin filaments (Fig. 4), resembling the distribution of α-actinin and palladin (39,40).
Myotilin stabilizes actin filaments
In order to investigate possible effects of myotilin on actin filament stability, we assayed the disassembly rate of pyrene-labelled actin filaments mixed with different amounts of His-tagged, full-length myotilin and 20 µm Latrunculin A (LatA), an actin monomer sequestering agent (45). The disassembly process was quantified using a fluorometer at an emission wavelength of 407 nm. Within 10 min, LatA significantly increased the depolymerization rate of F-actin (Fig. 5A, lowest curve). Upon addition of myotilin at a 1:4 molar ratio to actin, the disassembly rate decreased by 10%. With increasing concentrations of myotilin the rate of disassembly was further decreased and at a 1:1 actin:myotilin ratio, the rate decreased to 40% of that of F-actin alone.
The filament-stabilizing effect of myotilin was further studied in vivo in CHO-cells expressing transfected myotilin. In control cells, a 30 mm LatA treatment caused a pronounced dissociation of filamentous actin, and only patches of F-actin remained (Fig. 5B). In myotilin expressing cells, however, the myotilin-induced actin-bundles were resistant to the depolymerizing effect of LatA. These results suggest that binding of myotilin to actin filaments decreases the dissociation of actin monomers from filaments and thus protects the filaments from depolymerization.
Myotilin expression is initiated at the stage of myofibril alignment
In order to study the temporal expression pattern of myotilin in differentiating human skeletal muscle cells, RNA extracted from cultured cells differentiated for 1–6 days was used in northern blotting experiments. At the same time, myotilin expression was investigated at the protein level by western blotting using detergent soluble and insoluble fractions of cells lysed in Triton X-100 containing buffer. Figure 6 shows that the expression of myotilin mRNA starts at a low level already at the third day of differentiation (panel A). The expression level gradually increased and strong mRNA signal was detected at day 6 of differentiation. At the protein level, significant myotilin expression could only be detected at the latest stages of differentiation (day 6, panel B). In contrast, sarcomeric α-actinin was already expressed at detectable levels after the first day of differentiation (Fig. 6).
In line with our biochemical studies, immunofluorescence microscopy of differentiating cultured human skeletal muscle cells demonstrated barely visible myotilin staining at initial stages of myofibril assembly (Fig. 7A and B). Only at the time-point when myofibrils began to align laterally was a strong increase in myotilin staining intensity observed at the level of Z-discs (Fig. 7C and D).
In sections of mature, normal human skeletal muscle, staining with myotilin antibody revealed a cross-striated pattern (Fig. 7E) and double staining with an antibody recognizing a Z-disc epitope of titin confirmed myotilin's Z-disc localization (data not shown). Immunoelectron microscopy demonstrated decoration of the central Z-disc portion (Fig. 7F).
Truncated myotilin fragments exhibit a dominantnegative effect on myofibril assembly
We also wanted to investigate which portions of myotilin might be involved in myofibril assembly. In addition to the full-length cDNA, the amino-terminal (Myot 1–251) and carboxy-terminal (Myot 252–498), half of myotilin were cloned into the newly constructed pMYP-vector (see Material and Methods). In this vector, a cDNA is expressed under control of a myomesin promoter fragment, which assures more steady expression at a level typical for muscle proteins at a medium level from day 3 of differentiation onwards. Transfected C2C12 cells were allowed to differentiate for up to 6 days. Thus, the premature actin filament bundling as observed in non-muscle cells (see Fig. 4) was avoided. Recombinant proteins were detected by immunostaining with a T7-tag antibody and forming myofibrils were visualized by Z-disc specific titin staining. During early stages of differentiation, both the amino- (Fig. 8A and B) and the carboxy-terminal myotilin fragment (Fig. 8E and F) showed a diffuse or filamentous staining pattern with a concentration of the expressed polypeptides in nascent Z-discs (Z-bodies, Fig. 8A, B, E and F), and did not exhibit dominant negative effects on myofibrillogenesis. However, as differentiation proceeded, both recombinant polypeptides started to accumulate in aggregates together with titin (Fig. 8C, D, G and H), indicating a severe disruption of myofibril assembly. In contrast, these dominant negative effects were never seen upon expression of the full-length recombinant myotilin (Fig. 8I and J). These results indicate that the complete myotilin protein is required for sarcomere integrity, and that both terminal regions are independently involved in as yet unidentified control mechanisms of myofibril assembly.
In the present study we have shown that myotilin does not only bind α-actinin (27) and filamin c (38), but also F-actin. The direct association of myotilin with F-actin induces filament bundling both in vitro and in vivo. Furthermore, myotilin stabilizes the assembled actin bundles, probably by decreasing the off-rate of actin monomers from filament ends. The expression of myotilin mRNA starts relatively late during muscle cell differentiation and the protein does not appear at the Z-discs until the time of myofibril alignment. The expression of myotilin in non-muscle cells leads to a dramatic reorganization of the actin cytoskeleton. In contrast, expression of myotilin in differentiating muscle cells under control of the myomesin promoter, which is only activated around the third day of in vitro myocyte differentiation, results in proper incorporation of the recombinant protein into Z-discs and allows for normal myofibril organization. This indicates that an appropriate regulation of myotilin expression is important for the normal assembly of myofibrils. We further conclude that in developing myofibrils myotilin is functional at the developmental stage when the actin cytoskeleton has already transformed from a stress fibre-like arrangement with its long filaments to the short bipolar I-Z-I brushes. This view is supported by experiments in which the transfection of amino-terminal or carboxy-terminal deletion constructs results in the disassembly of nascent sarcomeres, and an accumulation of myofibrillar proteins in aggregates.
Myotilin binds and bundles actin filaments
Both full-length myotilin and a fragment comprising amino acids 80–462 directly bound F-actin. Furthermore, transfection studies with the carboxy-terminal construct Myot 215–498 in non-muscle cells showed a clear co-localization of the expressed protein and the cortical actin cytoskeleton suggesting that the actin-binding site resides in the Ig-like domain-containing part of myotilin. At the same time, the Ig-domain region of myotilin also seems to be involved in the self-association that is a prerequisite for the observed actin cross-linking activity. In ABP-120 and in the vertebrate filamins, an Ig-like domain can also form the interface for dimerization (4). The stability of cross-linked actin bundles is thought to be enhanced by high local concentrations of both cross-linkers and actin filaments in these assemblies (4). In addition, cross-linkers can simply impose a steric hindrance, which decreases the rate of F-actin disassembly. However, an F-actin cross-linking protein in Dictyostelium was shown to prevent F-actin depolymerization independently of the cross-linking event (46). Our Latrunculin A experiments demonstrate that myotilin can prevent actin filament depolymerization efficiently both in vivo and in vitro, suggesting that myotilin protects thin filaments in muscle cells from disassembly at their barbed ends. Furthermore, myotilin might have a role in the lateral alignment of nascent myofibrils, by integrating I-Z-I bodies, preformed arrays of thin filament and Z-disc proteins (28,36), into sarcomeres, and in the stabilization of the assembled Z-discs.
Myotilin is a component of a complex of multiple actin cross-linking proteins
A further excellent example where at least two cross-linking proteins are required for proper organization of actin filaments is the formation of bristles in Drosophila. The ‘forked’ protein is used early in the development to tie up actin filaments into tiny bundles. Subsequently, forked protein orchestrates fascin entry into the bundles, leading to cross-linked, straight, compact and rigid bundles (47). Forked protein is thought to keep the bundles in a partially completed state, enabling fascin to quickly find its place in the assembly and to strengthen the bundles with the second cross-links. This model is particularly attractive in light of the order of appearance of α-actinin and myotilin into forming Z-discs. However, in Z-discs the picture is even more complex, since myotilin is not only a ‘second cross-linker’ that integrates into a pre-existing structure, it also binds two other actin filament cross-linkers, α-actinin and filamin c, thus forming a unique complex of three actin cross-linkers at the myofibrillar Z-disc. Both, α-actinin and filamin c are among the first myofibrillar proteins to be expressed during differentiation and together they might orchestrate the entry of myotilin into its correct position at the Z-disc. In fact, the region around the Ig-domain containing the muscle specific insertion in filamin c, which seems to be involved in myotilin interaction, is already sufficient to relocate recombinant myotilin in double-transfected non-muscle cells (38). The capacity to induce bundling of actin filaments both in vitro and in vivo and the late expression of myotilin are therefore supporting a ‘fascin-type’ mode of action for myotilin. We propose that myotilin serves as an important link for thin filament anchorage at the Z-discs, connecting several Z-disc components (Fig. 9) and providing rigidity and strength to the complex such that it can resist mechanical stress upon muscle contraction.
Myotilin is functional during later stages of myofibril assembly
Endogenous expression of myotilin in skeletal muscle cells is strictly controlled and occurs only at the later stages of in vitro differentiation, which is in striking contrast to most other sarcomeric proteins. This exceptional temporal expression pattern makes myotilin a rather unique member of the I-band proteins. Since the overexpression of myotilin has a dramatic, one-of-a-kind impact on the organization of the actin cytoskeleton in non-muscle cells, we suppose that its delayed expression in skeletal muscle cells is required to overcome premature actin bundling that might hinder the normal assembly of myofibrils. With the construction of an expression vector containing the myomesin promoter (pMYP), which is active only at later stages of myofibrillogenesis (34,48–50), we avoided premature myotilin expression.
Transfection experiments were subsequently designed to test for the relative importance of distinct portions of myotilin for sarcomere integrity. We thus found that both amino- and carboxy-terminally truncated constructs in the pMYP vector initially bound to nascent myofibrils, but subsequently caused disruption of nascent myofibrils followed by an aggregation of myofibrillar proteins. A similar series of events was reported for transfection of α-actinin constructs lacking EF-hands in the carboxy-terminal portion (51). In chicken smooth muscle α-actinin the binding site for myotilin was reported to reside in its spectrin-like repeats (27). Recent experiments with mammalian sarcomeric α-actinin have located the myotilin-binding site to the carboxy-terminal EF-hand region (A.Taivainen et al., manuscript in preparation), which also harbours a binding site for titin (52,53). The fact that overexpression of truncated α-actinin and myotilin variants results in similar phenotypes hints at the important function of their intermolecular interactions for Z-disc assembly and stabilization. Although our emerging knowledge about the complexity of protein–protein interactions in this sarcomeric structure has increased rapidly during the last 5 years, the precise role of the individual constituents in myofibrillogenesis is still far from being understood.
The present study sheds new light on the pathogenesis of LGMD1A caused by myotilin mutations (42,43). Although the clinical picture in LGMD1A has common features with other forms of LGMD, most often resulting from a dysfunctional sarcoglycan complex, the morphological alterations are distinct and suggest differences in the molecular events that cause the phenotype of the disease. Prominent structural abnormalities in muscles affected by LGMD1A include extensive Z-line streaming and myofibril aggregation that are consistent with the idea that the primary defect involves proper maintenance of the sarcomeric structure. This idea is further supported by the finding that mutations in the Z-disc protein telethonin also cause LGMD (54). Our results, which identify myotilin as a key structural component of the Z-disc, will now enable more focused studies aimed at directly understanding why a single amino acid change (S55F, T571) in myotilin manifests as a clinical disease, and leads to the observed LGMD1A phenotype.
MATERIALS AND METHODS
Antibodies, purified proteins and cDNA constructs
The following primary antibodies were used in this work: mouse anti-HA mAb (12CA5) recognizing the HA-tag (Roche Diagnostics); mouse T7-tag mAb (IgG2b, Novagen); mouse T12 mAb (IgG1) directed against an epitope located at the Z-disc region of titin (55); RaA653, a rabbit polyclonal antiserum specific for skeletal muscle α-actinin (22); mouse mAb EA53 (IgG1), directed against sarcomeric α-actinin (56); and rabbit polyclonal antibody 948, directed against myotilin (27). As secondary antibodies, FITC-conjugated goat anti-mouse IgG (Cappel Research Products), TRITC-conjugated goat anti-rabbit F(ab)2 fragment (Jackson Immunoresearch Laboratories), FITC-conjugated anti-mouse IgG1 (Rockland Inc.) and TRITC-conjugated anti-mouse IgG2b (Rockland Inc.) were used. Purified rabbit muscle actin (AKL 99), pyrene-labelled actin (AP04) and rabbit skeletal muscle α-actinin (AT01) were purchased from Cytoskeleton Inc. The coding region of myotilin (full-length, wt) and partial fragments 1–251, 116–498, 215–498, 252–498 and 80–462 were amplified from the original library clone (27). PCR products were digested with restriction enzymes and ligated into pAHP, pGEX, pKKtac and pMYP vectors. For the construction of pMYP, a part of the myomesin promoter (bp −596/+122) that was shown to be active only in differentiating muscle cells (57) was cloned in the pCAT3E vector in its multiple cloning site upstream of the CAT cDNA (Promega). Subsequently the cDNA encoding CAT was replaced by a novel multiple cloning cassette containing unique restriction sites and a sequence encoding a carboxy-terminal T7-tag. The authenticity of all constructs was verified by sequencing. Plasmid DNAs were purified using the Maxiprep kit (Qiagen). The human non-muscle α-actinin-pEGFP-N3 construct was a kind gift from Dr. Carol Otey (University of North Carolina, Chapel Hill, NC, USA) and the α-actinin R1–R4-pET construct from Drs Paul Young and Mathias Gautel (Max Planck Institute for Molecular Physiology, Dortmund, Germany).
Production of recombinant proteins
Full-length myotilin cDNA and the fragment 116–498 in pKKtac vector were transformed into E.coli BL21 (pLys-S). Protein expression was induced with 0.5 mm IPTG for 2 h at room temperature and the His-tagged fusion proteins were purified with Ni-Sepharose. Immediately after elution, the buffer of purified proteins was changed into 20 mm Tris, 50 mm NaCl, pH 8 using PD10-columns (Amersham Pharmacia Biotech). Another myotilin construct (amino acids 80–462) in pGEX 4T-1 vector was transformed into E.coli BL21 (DE3) cells, induced with 0.4 mm IPTG and purified on glutathione-Sepharose beads (Amersham Pharmacia Biotech). The buffer of this GST-fusion protein was also changed into 20 mm Tris, 50 mm NaCl, pH 8 after purification. α-actinin fragment R1–R4 was produced and purified as described (52).
Actin filament co-sedimentation and bundling assays
For cosedimentation and bundling assays, actin filaments were assembled from purified rabbit skeletal muscle actin in G-buffer (5 mm Tris, pH 8, 0.2 mm CaCl, 0.5 mm DTT, 0.2 mm ATP) by addition of 0.1×volume of 10× polymerizing mixture (500 mm KCl, 20 mm MgCl2, 10 mm ATP) and incubated at 25°C for 30 min. In the cosedimentation assay, GST-myotilin (1.5 µm) or purified α-actinin (2 µm) was added to the preassembled actin (7 µm) filaments. In the bundling assay, His-myotilin (1.5 µm), α-actinin (1 µm) and/or α-actinin R1–R4 (1 µm) was added to actin (3 µm) filaments. Binding stochiometry was evaluated by adding variable concentrations (0.12–2 µm) of His-myotilin to premade actin (1 µm) filaments. Reaction volumes were equalized to 50 µl with G-buffer. Mixtures were incubated at 25°C for 30 min and subsequently centrifuged as follows: for co-sedimentation, 10 000g, 1 h in a Beckman TL-100 ultracentrifuge; for bundling assays, 14 000g, 3 min in an Eppendorf centrifuge. Pellets and supernatants were separated and their volumes were equalized with SDS sample buffer. Samples were subjected to SDS–polyacrylamide gel electrophoresis and proteins were visualized by Coomassie brilliant blue staining and quantitated with a densitometer.
Actin filament stabilization assay
For fluorometric measurement of F-actin disassembly, purified, pyrene-labelled actin and unlabelled actin were diluted in G-buffer. In 50 µl reactions, 4 µm actin (2 µm actin+2 µm pyrene-actin) was preassembled to filaments, and variable amounts of His-tagged myotilin were added. After 30 min incubation, 2 µl of 500 µm Latrunculin A (Calbiochem) was mixed with each reaction and the mixture was immediately transferred to a quartz glass fluorometer cuvette with 3 mm light path (Helma). Actin filament disassembly was monitored by pyrene fluorescence with excitation at 365 nm and emission at 407 nm in a fluorescence spectrophotometer (Hitachi).
Human myotilin fragment aa 215–442 was cloned into PGEX-4T-I and the construct was transformed into E.coli BL21 (pLys-S). Protein expression was induced with 0.2 mm IPTG overnight at 18°C. Bacteria were processed at 4°C using a French Pressure Cell Press (SIM Aminco). The fraction that remained in the supernatant after centrifugation at 30 000g was applied to gluthathione Sepharose 4B beads (Pharmacia Biotech) for 3 h (4°C) and the fusion protein was cleaved using 10 U of Thrombin (Sigma) in 50 mm Tris–HCl, pH 8.0, 150 mm NaCl, 2.5 mm CaCl2, for 2 h at RT. Subsequently, beads were washed four times in the same buffer, and thrombin was removed by benzamidine Sepharose chromatography. After buffer exchange to 50 mm Hepes, pH 7.5 using PD-10 columns, the protein was applied to a POROS HS-M cathione exchange column (4.60/100 mml=1.662 ml, Biocad Aplied Biosystems) and eluted by gradient of 1 m NaCl in the same buffer. Following elution, the protein was concentrated in 20 mm Tris, pH 8.5, to a final concentration of 0.1–0.4 mm.
Fractions of 100 µl were analyzed by size exclusion chromatography on Superdex 10/30 HR column in 20 mm Tris–HCI, pH 8.5, at 8°C by using the SMART system (Pharmacia Biotech). Absorbance was measured at 280 nm. Flow rate was set at 0.3 ml/min and 900 µl fractions were collected. The resulting fragment (25 kDa) was subjected to N-terminal sequence analysis on a 477A protein sequencer with an on-line 120A PTH analyser (Applied Biosystems).
For MALDI-TOF (matrix-assisted laser desorption/ionization time of flight mass spectrometry), SDS–PAGE and Western blotting, fractions of the sample were concentrated using reverse-phase chromatography C4 zip-tips (Millipore) and eluted with 100% acetonitrile. For matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) samples were dried down to ∼5 µl. One microliter of the product was mixed with Sinapinnic acid (CA), allowed to cocrystallize and analyzed further on Biflex III (Bruker) instrument in linear mode. All spectra were externally calibrated using a lock mass routine based on monoistopic molecular ions relating to α- and ß-lactoglobulin product presents at 18278.4 and 18364.3 Da.
COS-1- and CHO-cells (ATCC) were grown in DME medium supplemented with 10% FCS, 1% l-glutamine, 100 U/ml penicillin and 1 µg/ml streptomycin. Cells were seeded on glass coverslips one day before transfection and grown to ∼30–40% confluency. Culture medium for C2C12 cells (no. CB2438, ECACC) was DME supplemented with 15% FCS and antibiotics. Differentiation was induced by transferring cells to a low nutrient medium (2% horse serum instead of FCS). C2C12 cells were seeded on glass coverslips 6 h prior to transfection and grown to ∼80% confluency. All cells were transfected with Fugene6 (Roche Diagnostics, Indianapolis) according to manufacturer's instructions, except that with C2C12 cells, 5 µg of plasmid DNA and 6 µl of Fugene were used per 35 mm well. Transfected C2C12 cells were allowed to differentiate by changing the culture medium to the low nutrient medium one day after transfection.
mRNA and protein studies
Human skeletal muscle cells were isolated and cultured essentially as described (38). Frozen cells from liquid nitrogen were quickly thawed, plated and grown in DME medium supplemented with 20% FCS, 2% Ultroser G, 100 U/mI penicillin and 1 µg/µl of streptomycin (all from Life Technologies). Differentiation was induced by changing the culture medium to low-nutrient differentiation medium (0.4% Ultroser G in DME). Two sets of cells were grown on 60 mm plates and differentiated for 0–6 days. One set was subjected to mRNA isolation (Qiagen) and subsequent Northern blotting. Five micrograms of each mRNA were mixed with RNA loading buffer (50% glycerol, 1 mm EDTA, 0.25% Bromphenolblue, 0.25% xylene cyanol) and incubated at 65°C for 15 min. Samples were run in a formaldehyde-agarose gel and transferred to a nitrocellulose membrane (Schleicher & Schuell). 32P-labelled full-length myotilin cDNA and rat GAPDH cDNA were used as probes in the subsequent hybridization. The other set of cells was extracted with Triton-buffer (1% Triton X-100, 50 mm NaCl, 300 mm sucrose, 10mm Pipes, 3 mm MgCl2, pH 6.8 and protease inhibitors) to fractionate detergent soluble and insoluble components. Equalized volumes of both fractions were boiled in SDS sample buffer and proteins were separated on an 8% polyacrylamide gel. Proteins were transferred to nitrocellulose filters using a semi-dry blotting apparatus (BioRad) and the blots were incubated with antibodies specific for sarcomeric α-actinin or myotilin, and horseradisch peroxidase-conjugated secondary antibodies. Immunoreactivity was visualized using the ECL detection method (Pierce).
Actin filaments from the bundling assay (without centrifugation) were directly applied to glow-discharged formvar-carbon coated nickel grids. After 5 min, the liquid was removed and filaments were negatively stained with aqueous 1% uranyl acetate. The grids were air-dried and observed in a JEOL 1200 EX transmission electron microscope operated at an accelerating voltage of 80 kV.
For immunoelectron microscopy normal human skeletal muscle tissue was fixed in a solution of 4% paraformaldehyde and 0.1% glutaraldehyde with HEPES buffer (0.1 m, pH 7.5), washed for 3×min with the same buffer, dehydrated with series of ethanol and embedded in LR White®. Ultrathin sections were rinsed on drops of washing buffer for 10 min, etched for 15 min with saturated sodium metaperiodate, followed by a rinse in water and 10 min in 0.1 m HCL. After etching, grids were rinsed with water for 10 min and preincubated in 5% normal goat serum in washing buffer (PBS, 0.5% BSA, 0.1% fish gelatin) for 35 min at room temperature. The grids were washed for 10 min with washing buffer and then incubated with the primary desmin antibody diluted 1:20 in washing buffer (with 1% normal goat serum) in a moist chamber at 4°C. After overnight incubation with the primary antibody the grid was washed with washing buffer and incubated with secondary antibody (Aurion) coupled to colloidal gold (diluted 1:20; 10 nm particles). The grids were washed with washing buffer (4 × 3 min), followed by four washes in PBS and postfixed with 1% glutaraldehyde in PBS. Following another wash with distilled water (2 × 5 min), the preparations were contrasted using conventional techniques. Ultrathin sections were examined in a Zeiss 900 electron microscope at an accelerating voltage of 80 kV.
Mrs Helena Ahola, Mrs Maija-Liisa Mäntylä and Mrs Bärbel Mai are gratefully acknowledged for their skilful technical assistance, Carol Otey, Paul Young and Mathias Gautel for the α-actinin constructs and Marc Baumann for advice and help in dimerization studies. This study was supported by Academy of Finland, Sigrid Juselius Foundation, TEKES, the Cancer Society of Finland, the Finnish Muscular Disease Research Foundation, the German Research Foundation, and the BMBF (Bundesministerium für Bildung und Forschung).
To whom correspondence should be addressed at: Department of Pathology and Neuroscience Program, University of Helsinki, Biomedicum, PO Box 63, 00014 Helsinki, Finland. Tel: +358 919125650; Fax: +358 947171964; Email: firstname.lastname@example.org