The M band of sarcomeric muscle is a highly complex structure which contributes to the maintenance of the regular lattice of thick filaments. We propose that the spatial coordination of this assembly is regulated by specific interactions of myosin filaments, the M band protein myomesin and the large carboxy‐terminal region of titin. Corresponding binding sites between these proteins were identified. Myomesin binds myosin in the central region of light meromyosin (LMM, myosin residues 1506–1674) by its unique amino‐terminal domain My1. A single titin immunoglobulin domain, m4, interacts with a myomesin fragment spanning domains My4–My6. This interaction is regulated by phosphorylation of Ser482 in the linker between myomesin domains My4 and My5. Myomesin phosphorylation at this site by cAMP‐dependent kinase and similar or identical activities in muscle extracts block the association with titin. We propose that this demonstration of a phosphorylation‐controlled interaction in the sarcomeric cytoskeleton is of potential relevance for sarcomere formation and/or turnover. It also reveals how binding affinities of modular proteins can be regulated by modifications of inter‐domain linkers.
The impressively regular and stable organization of thick and thin filament lattices in the sarcomeres of cross‐striated muscles cannot be explained by the self‐assembly properties of their major constituents, actin and myosin. Instead, it is brought about by a cytoskeletal framework whose complexity was only appreciated recently (see, e.g., review by Small et al., 1992). The most obvious structures involved in the regular packing of contractile filaments are the Z discs and M bands, which organize thin and thick filaments, respectively. The only component identified to date that could integrate both structures is the giant protein titin (also called connectin) together with its associated proteins (for a recent review, see Fürst and Gautel, 1995).
A small group of proteins specifically located in the M band is known. Apart from the ATP‐regenerating enzyme MM‐creatine kinase, three structural proteins have been established in this region: M‐protein (Masaki and Takaiti, 1974), myomesin (Grove et al., 1984) and titin (Nave et al., 1989). M‐protein originally was characterized biochemically as a myosin‐associated protein (Masaki and Takaiti, 1974; Trinick and Lowey, 1977) and myomesin was identified as a contaminant in M‐protein preparations by using monoclonal antibodies (mAbs) (Grove et al., 1984). The function of both proteins remained largely enigmatic. The finding that both myomesin and M‐protein also bind to titin suggests that they have essentially cytoskeletal functions (Nave et al., 1989; Vinkemeier et al., 1993). While myomesin is found in all striated muscle fibres examined, M‐protein is restricted to fast skeletal and cardiac fibres (Grove et al., 1985, 1987).
A structural model describing the exact disposition of titin, myomesin and M‐protein in the M band was only introduced recently (Obermann et al., 1996). The availability of cDNA information for titin, myomesin and M‐protein enabled the production of a panel of specific and sequence‐assigned antibodies and the subsequent localization of the respective epitopes at the sarcomere level by electron microscopy. In this model, the carboxy‐terminal end of titin spans the central M1 line and reaches ∼60 nm into the other sarcomere half. While myomesin was also implied to bridge the M1 line and to be arranged largely in an antiparallel and staggered fashion, M‐protein could bridge the thick filaments primarily in a perpendicular orientation (Obermann et al., 1996). This model puts certain constraints on protein–protein interactions necessary to achieve the three‐dimensional structure of the M band. We approached this problem by producing a series of recombinant fragments of myomesin, of the LMM portion of myosin and of the 250 kDa carboxy‐terminal region of titin, and have used these to search for mutual binding sites. The results are in good agreement with the previously proposed model. The possibility of regulating at least one of the binding sites by phosphorylation invites some speculation on its function during sarcomere formation and/or turnover.
A novel phosphorylation site in the loop between domains My4 and My5 of myomesin
In the myomesin molecule, a unique amino‐terminal domain (138 residues) is followed by 12 repeat domains which reflect immunoglobulin cII and fibronectin type III domains respectively (Vinkemeier et al., 1993; Figure 1). Although computer analyses of the human sequence did not indicate specific target sites for protein kinases (Vinkemeier et al., 1993), we observed that the phosphorylation‐dependent mAb NE‐14, which is directed against porcine neurofilament H protein (Shaw et al., 1984), shows cross‐reactivity with purified bovine myomesin. The corresponding phosphorylation site locates to the ∼60 carboxy‐terminal residues of myomesin (Obermann et al., 1995). To search for kinases acting on myomesin, the purified protein was incubated with three distinct protein kinases in the presence of [32P]ATP. While cAMP‐dependent protein kinase A (PKA) was able to phosphorylate myomesin (Figure 2A), protein kinase C and casein kinase showed no reaction (data not shown). For a preliminary mapping of the phosphorylation site, we used the limited proteolysis procedure with endoproteinase Lys‐C which generates two myomesin fragments with Mrs of 116 and 109 kDa, respectively. Since both fragments were shown by direct protein sequencing to start with Ser479, their mass difference must reside in a carboxy‐terminal end that is ∼60 residues shorter in the lower molecular mass fragment (Obermann et al., 1995). If, as we assumed, the PKA phosphorylation site was identical to the site previously identified in this region by mAb NE‐14, only the higher molecular mass band should be phosphorylated. Figure 2A (lane 2) shows an autoradiograph of the Lys‐C limited digest of bovine myomesin previously radioactively phosphorylated by PKA. Surprisingly both fragments are radioactively labelled. Thus, myomesin shows an unexpected phosphorylation site which must lie between Ser479 and the last residue common to the two fragments.
For a more precise mapping of the PKA phosphorylation site, we used a series of overlapping myomesin constructs amplified from the human cDNA by PCR and cloned into the modified bacterial expression vector pET described before (Obermann et al., 1996). Corresponding recombinant proteins were overexpressed in Escherichia coli and purified under native conditions making use of their carboxy‐terminal (His)6 tag to facilitate purification. Figure 2B shows the purity of the recombinant myomesin fragments used in the phosphorylation studies and their location with respect to the domain organization of myomesin is given in Figure 1. Recombinant fragments were treated with PKA and ATP under standard assay conditions. SDS–PAGE and corresponding autoradiographs (Figure 2B and C) showed that myomesin My4–5 and larger fragments containing these two domains (see My4–8) are phosphorylated efficiently, while My1–3, My6–8 and My9–13 are not substrates for PKA. Phosphoamino acid analysis of radiolabelled My4–5 and myomesin showed that PKA phosphorylation occurs exclusively on serine (Figure 3A). Closer inspection of the sequence of My4–5 in the human myomesin sequence (Vinkemeier et al., 1993) hinted at the rather long insertion connecting these two FN domains (see also Figure 1). This region of ∼35 residues harbours a sequence containing four basic residues, two serines and two prolines, KARLKSRPSAP (residues 474–484), which is more likely to be accessible than the serines in the tightly packed β‐barrels of the repeat domains. Therefore, a synthetic peptide EKARLKSRPSAPWTGQ (residues 473–488) covering this region was tried as a PKA substrate. This peptide is phosphorylated efficiently by PKA. Since this sequence describes a novel target site for PKA, the kinetics of the phosphorylation reaction were compared with the kinetics of the liver pyruvate kinase peptide LRRASLG (‘kemptide’), which is an excellent substrate for PKA with a Km of 16 μM and a Vmax of 20.2 μmol/min/mg (Kemp et al., 1977). Table I shows that the apparent Km value of kemptide measured under our assay conditions was nearly identical to the published value. The Km of the myomesin peptide EKARLKSRPSAPWTGQ was 532 μM. The Vmax was not obtained in absolute numbers, since the enzyme concentration in the commercially available sample of PKA and in muscle extracts could not be measured. Instead, the numbers of our measurements in Table I indicate relative units. Thus the Vmax of the myomesin peptide phosphorylation reaction is ∼50% of that of the kemptide reaction. Reverse phase HPLC (Figure 3B) shows a clear shift of the elution volume of the peptide after phosphorylation. This mobility shift indicates that under standard assay conditions the peptide is converted quantitatively into the phosphorylated form. Phosphorylation of the peptide was, as in My4–5, serine specific (data not shown).
To decide which serine in residues 474–484 is the target site for PKA, two constructs comprising myomesin My4–5 were made and either serine 479 or 482 was substituted by alanine. The constructs were expressed in E.coli and the two recombinant myomesin fragments carrying a single point mutation were purified in the native state. Subsequent phosphorylation assays with PKA monitored by SDS–PAGE and autoradiography (Figure 2B and C) demonstrated that My4–5 (Ser479/Ala) is still phosphorylated while My4–5 (Ser482/Ala) is no longer a substrate of PKA. Thus myomesin Ser482 is the phosphorylation site for PKA.
Myomesin is phosphorylated by skeletal muscle extracts
A series of extracts from muscle tissues was tested for their ability to incorporate radioactive phosphate into myomesin. The results resembled the in vitro observations with PKA. Native myomesin purified from bovine skeletal muscle and the recombinant fragments listed in Figure 2 were phosphorylated as described above for purified PKA (data not shown). No difference was observed with respect to the state of differentiation of the muscle tissue from which the extract was derived. Thus the degree of phosphorylation of the myomesin My4–5 fragment was the same with extracts from rat fetal (gestation day 14), neonatal and adult rat psoas muscle (results not shown). The ability of these extracts to phosphorylate Ser482 in the My4–5 fragment was also characterized in the presence of the PKA substrate kemptide (Kemp et al., 1977). It resulted in a complete inhibition of the phosphorylation of the myomesin fragment. Similarly, the presence of the regulatory subunit of PKA also led to complete inhibition of the phosphorylation of the myomesin fragment (results not shown). In contrast, addition of EGTA or Ca2+/calmodulin to phosphorylation assays did not affect incorporation of radioactive phosphate into myomesin My4–5. We conclude, therefore, that myomesin phosphorylation by muscle extracts is probably the result of PKA or an enzyme closely related to PKA.
Interaction of myomesin with titin is modulated by phosphorylation of Ser482
Overlapping myomesin fragments spanning the entire molecule and nine domains from the carboxy‐terminal (M band) region of titin (see scheme in Figure 1) were expressed in E.coli and purified by Ni–NTA chelate affinity chromatography followed by ion‐exchange chromatography. Figure 4 documents the purity of the panels of purified myomesin fragments (My1–8, My1–5, My4–5, My4–8, My5–6, My9–13, My4–6 and My7–8) and titin fragments (m1, m2–m3, m4, m5–m6, m7, m8, m8–m9, m9, m10 and the titin kinase domain) by SDS–PAGE. A solid phase overlay assay was used to investigate the myomesin–titin interaction (Figure 5). Interestingly, m4 is the only titin domain which shows highly selective interaction with myomesin fragments My1–8 and My4–8. Since the titin fragment m5–m6, which lies next to m4 (see domain structure in Figure 1) lacked myomesin binding (Figure 5), we also used m5–m6 after phosphorylation by cdc2 kinase in the overlay assay. Again, no binding of myomesin fragments was observed (Figure 5).
For a more precise mapping of the titin m4 binding site within the five domains of myomesin fragment My4–8, a series of constructs representing different portions of this region were made. Surprisingly, all of the myomesin fragments comprising only two FN domains lost the ability to interact with the titin domain m4 (Figure 5; for domain structure see Figure 1). In contrast, myomesin fragment My4–6, which covers the first three FN domains, retained binding to titin (Figure 5). The results were the same whether titin domains were spotted onto nitrocellulose and overlayed with myomesin fragments, or the reverse order was used. This demonstrates the selectivity and specificity of the solid phase overlay assay used.
Since myomesin fragment My4–6 contains the PKA phosphorylation site identified above, it was particularly interesting to see whether phosphorylation of Ser482 would influence the binding to titin m4. Therefore, the binding assay was repeated using the myomesin fragment My4–6 either directly or after PKA phosphorylation (Figure 5B). Interestingly, phosphorylation almost completely abolished the binding of My4–6 to titin domain m4.
The amino‐terminal head domain of myomesin binds to myosin; location of the myomesin binding site on the myosin rod
A previous study reported binding of myomesin to the LMM portion of myosin (Obermann et al., 1995). For a more detailed mapping of the binding sites in both proteins, a series of recombinant myomesin fragments, and several proteolytic and recombinant derivatives of myosin were made. Figure 6A and B shows the purity of these derivatives. Again a solid phase overlay assay was used to delineate the sites of interaction. While myomesin fragments My2–3, My4–8 and My9–13 lacked binding to LMM, the myomesin fragment My1–3, which contains the head domain, showed a concentration‐dependent binding to LMM (Figure 6C). This finding suggests that the unique amino‐terminal head portion of the myomesin molecule bears the myosin binding site. The location of this binding site within the myomesin molecule is also indicated in the schematic representation given in Figure 7.
In the same way, different myosin derivatives were used to map the myomesin binding site on the rod portion of myosin with greater precision. While proteolytic myosin rod, proteolytic and recombinant LMM as well as LMM 59 and LMM 50 were recognized by the myomesin fragment My1–3, LMM 50–75 and LMM 30 failed to bind myomesin My1–3 (Figure 6C). We conclude, therefore, that the myosin binding site on the myosin heavy chain is confined to the 169 amino acids between residues 1506 and 1674 located in the central part of LMM (numbers refer to the amino acid sequence of rabbit fast skeletal muscle myosin heavy chain; see Maeda et al., 1987).
As a ubiquitous protein of vertebrate sarcomeric M bands, myomesin can be supposed to play a key role in establishing this complex structure. At the level of isolated proteins, it has been established that myomesin binds to both titin (Nave et al., 1989) and myosin (Obermann et al., 1995). Thus, myomesin may act as a cross‐linker connecting the M band end of titin with myosin thick filaments. It is important, therefore, to map the respective binding sites in detail and to search for possible regulatory mechanisms.
Our recent biochemical characterization of myomesin purified from bovine skeletal muscle revealed phosphorylation present within the carboxy‐terminal ∼60 residues of the polypeptide (Obermann et al., 1995). In an effort to search for the corresponding protein kinase, we now found a second, novel phosphorylation site. The combination of several approaches clearly established that Ser482 is the only residue that is phosphorylated specifically in vitro by PKA (Figures 2 and 3). This serine lies in the region connecting myomesin domains My4 and My5 (Figure 1) and is situated in the sequence KARLKSRPS* AP (residues 474–484; Vinkemeier et al., 1993). While it is more difficult to envisage that a particular sequence located within any of the densely packed Ig or FN repeat domains of myomesin would be a likely kinase target site, most of the basic residues in the loop sequence can be expected to be exposed to the surrounding medium and therefore to be a part of a kinase site. This particular site in myomesin also describes a novel target sequence for PKA whose preferred targets show two basic residues separated by one residue from a serine or threonine (Pearson and Kemp, 1991). In the myomesin sequence, four lysine/arginine residues provide a basic environment amino‐terminal to the phosphorylatable serine. Correspondingly, we find that a synthetic peptide spanning residues 473–488 is phosphorylated quantitatively by PKA under standard conditions. The apparent Km and Vmax values of this reaction (see Table I) indicate that myomesin is indeed capable of serving as a PKA target comparable with other substrates (Pearson and Kemp, 1991). When these values are compared, one has to take into account that most of the substrates characterized so far are soluble proteins. While for these proteins low Km values are essential to enable efficient phosphorylation in the cytosol, the high local concentration of myomesin in the M band may allow for higher Km values.
The possible significance of this phosphorylation site for the in vivo situation during myofibrillogenesis is emphasized by the observation that extracts of various muscle tissues of different developmental stages (from gestation day 16 rat embryos to adult) exhibit similar kinase activities on purified myomesin or its fragment My4–5 which harbours Ser482. It remains to be seen whether PKA phosphorylation of myomesin temporarily suppresses myomesin interactions during the early stages of myofibrillogenesis in vivo.
In this context, the localization of a titin binding site in the myomesin molecule established in this study is particularly interesting. Only fragments containing at least the three FN domains My4–6 exhibited binding to the titin Ig domain m4 (Figure 5). Single domains or pairs of neighbouring domains from My4–5 were not reactive in the solid phase overlay assay. This observation seems to resemble the situation found in the interaction between titin and cardiac C‐protein, where also a construct consisting of at least three C‐protein domains was identified as the minimal requirement for titin binding (Freiburg and Gautel, 1996). It appears that a larger number of weak but cooperative interactions is necessary to establish the stable complexes of titin with its various associated proteins that can be observed in vivo. It is probably essential for the ordered sequence of events during sarcomeric contraction–relaxation cycles that stable connections between contractile and cytoskeletal elements are retained. Most intriguing in this context is the finding that phosphorylation of a single serine residue (Ser482) within the myomesin binding site for titin resulted in an almost complete inhibition of binding (Figure 5B). Apparently the phosphorylation state of Ser482, which is located in the loop connecting myomesin domains My4 and My5, regulates the three‐dimensional arrangement of the three titin binding domains (My4–6) in a rather complex manner. The in vitro results invite some speculation on the molecular mechanisms of sarcomere formation, regeneration and turnover. In all cases, local signals for either the formation or the breakdown of certain complexes are needed. These could be provided by the phosphorylation/dephosphorylation of myomesin which in turn regulate its binding to titin. Likewise, the formation of stable links between titin molecules from neighbouring half sarcomeres might stabilize the cytoskeletal structures during myofibrillogenesis. The physiological role of myomesin phosphorylation in myofibril assembly remains now to be explored in the cellular context.
Solid phase overlay assays also delineated the molecular region of myomesin involved in the previously detected binding of myosin (Obermann et al., 1995). Since only constructs that contained at least myomesin domain My1 (i.e. the unique amino‐terminal domain) exhibited binding (Figure 6), we conclude that this domain either comprises the myosin binding site per se or is an essential part of a larger binding site. Since we could not stably express the My1 domain alone, we presently cannot distinguish between these two possibilities. Finally, we were able to locate the myomesin binding site on the myosin rod to residues 1506–1674 of the myosin heavy chain (Figure 6C; see Results). For a schematic representation of the established binding sites along the myomesin molecule, see Figure 7A.
The binding data presented here are in good agreement with the structural model of the sarcomeric M band region proposed on the basis of immunoelectron microscopical localizations of defined epitopes of myomesin, M‐protein and the carboxy‐terminal 250 kDa region of titin (Obermann et al., 1996; adopted in Figure 7B). The latter results have, for instance, suggested a particular arrangement of myomesin. While most of the myomesin domains seem organized parallel to the long axis of myofibrils, the amino‐terminal domains My1–2 appear to bend towards the thick filament. This region, most likely My1, indeed contains a myosin binding site (see above). Likewise, the binding of myomesin domains My4–6 to titin domain m4 described above requires close proximity of these regions to allow for these interactions to occur. This proximity is given in the structural model of the M band based on immunoelectron microscopy of defined epitopes of myomesin and titin. Here titin m4 occurs at a distance of ∼20–25 nm from the central M1 line, while the myomesin My4–6 region locates to a distance of ∼18–25 nm from the M1 line (see Figure 7B).
Materials and methods
Expression of titin M line domains and myomesin fragments in E.coli
Original λ phage isolates containing cDNAs coding for the M band portion of human cardiac titin (Gautel et al., 1993) were used as templates for PCR amplification (Saiki et al., 1985) of domains (see below). PCR products were cloned into a modified pET23a vector (Novagen, Heidelberg, Germany), providing the resulting proteins with an EEF tag at their carboxy‐termini. Since this tag is recognized by mAb YL1/2 (Wehland et al., 1984), expression in E.coli BL21(DE3)pLysS was monitored by immunoblot analysis. Purification of the soluble recombinant proteins on Ni–NTA–agarose columns due to their oligohistidine tag followed standard protocols (Qiagen, Hilden, Germany).
Myomesin sequences were amplified by PCR (Saiki et al., 1985) using the original λ phage isolates as templates (Vinkemeier et al., 1993). PCR products were cloned into pET‐23a derivatives. This provided the recombinant protein fragments with a carboxy‐terminally located His6 sequence and with either an amino‐terminally located T7 tag or a carboxy‐terminal EEF tag. After growth (LB medium supplemented with 2% glucose, 100 mg/l ampicillin and 34 mg/l chloramphenicol) and induction of E.coli BL21(DE3) LysS cells (Studier et al., 1990) at OD600 = 1 with 0.1 mM IPTG at 25°C for 3 h, the cells were harvested and stored at −70°C. Pellets were resuspended and sonicated in buffer A (50 mM potassium phosphate pH 8.0, 500 mM KCl, 0.02% Tween‐20, 5 mM 2‐mercaptoethanol) containing 5 μM E64 and 1 mM phenylmethylsulphonyl fluoride (PMSF) as protease inhibitors. After centrifugation at 16 000 g, soluble recombinant proteins were enriched by metal chelate affinity chromatography. Briefly, protein solutions were applied onto Ni–NTA–agarose columns (Qiagen, Hilden, Germany), which were washed with buffer A and subsequently with buffer B (same as buffer A, except pH 6.0). Finally the recombinant proteins were eluted with 500 mM imidazole in buffer B. After dialysis against buffer C [50 mM Tris–HCl, pH 7.9, 1 mM EDTA, 1 mM dithiothreitol (DTT)], recombinant proteins were purified further by ion‐exchange chromatography on MonoQ or MonoS FPLC columns (Pharmacia, Uppsala, Sweden). The integrity of the purified proteins was monitored by N‐terminal sequencing, the reaction with specific antibodies and the EEF tag‐specific antibody YL1/2 (Wehland et al., 1984).
Serine to alanine mutations were introduced into the myomesin My4–5 fragment by PCR amplification with mismatch oligonucleotides resulting in site‐directed exchanges of serine to alanine codons following the method of Ausubel et al. (1987). Amplified fragments were subcloned into pET‐23a‐derived expression vectors (see above). DNA sequencing established the original sequence and the desired Ser–Ala exchanges.
Recombinant fragments of myosin
Different portions of rabbit fast skeletal muscle LMM were amplified by PCR using the original cDNA of rabbit myosin heavy chain‐2/10 (Maeda et al., 1987) as a template. Four constructs were designed, in which the carboxy‐termini were identical with the end of the original clone (i.e. amino acid residue 1939). The amino‐termini were situated at different points along the sequence of the myosin rod portion. This resulted in a series of ‘shorter LMMs’, which were designated according to their approximate molecular masses (in kDa): LMM75 (corresponding to residues 1284–1939 of the original clone; Maeda et al., 1987), LMM59 (residues 1429–1939), LMM50 (residues 1506–1939) and LMM30 (residues 1675–1939). In addition, a further construct, called LMM75–50, was made. It corresponds to the amino‐terminal portion of LMM and represents residues 1284–1505 of the myosin heavy chain sequence. PCR products were cloned into the pET‐23a expression vector (Novagen, Heidelberg, Germany). Recombinant LMM fragments were expressed in E.coli BL21(DE3)pLysS and purified by two high salt–low salt extraction and precipitation cycles as described (Maeda et al., 1989). All recombinant LMM fragments formed normal paracrystals as judged by electron microscopic inspection after negative staining (Maeda et al., 1989).
Preparation of native proteins from bovine skeletal muscle
Myosin rod and LMM were prepared from bovine skeletal muscle by proteolytic cleavage with α‐chymotrypsin as described by Margossian and Lowey (1982). Purification of bovine skeletal muscle myomesin and its proteolytic fragments obtained by endoproteinase Lys‐C and trypsin have been described (Obermann et al., 1995).
Phosphorylation of proteins and synthetic peptides
Purified myomesin, its proteolytic fragments and recombinant myomesin fragments were added at a final concentration of 0.2 μg/μl to 20 μl of assay buffer (50 mM MES pH 6.9, 100 mM KCl, 2 mM MgCl2). Phosphorylation reactions were at 30°C for 30 min with 1 U of protein kinase A from porcine heart (Sigma) or 1 μl of muscle extract and 1 μCi of [γ‐32P]ATP (3000 Ci/mmol, Amersham). After addition of sample buffer (Laemmli, 1970) and heating at 65°C for 10 min, polypeptides were analysed by 4–12 or 6–20% SDS–PAGE. Autoradiography was at −80°C with intensifying screens.
For the in vitro phosphorylation of titin M5–M6, recombinant titin fragment M5–M6, containing the KSP repeats of the M band region of titin (Gautel et al., 1993), was expressed solubly and purified as described above. The purified protein was incubated in 100 μl of assay buffer (25 mM HEPES pH 7.2, 10 mM MgCl2, 1 mM EGTA, 1 mM DTT, 0.2 mM ATP) at 37°C for 30 min with recombinant cdc2‐ (New England Biolabs) or ERK‐2 (Stratagene) SP‐directed protein kinases, containing final concentrations of 0.1 mg/ml of the expressed substrate protein. Maximally, 2 mol/mol of phosphate could be incorporated as judged by parallel assays labelled with [γ‐32P]ATP and quantitation by liquid scintillation as described by Gautel et al. (1993). This indicates that in the presence of the flanking Ig domains, not all previously described SP phosphorylation sites are fully accessible.
Synthetic peptides were phosphorylated for kinetic experiments essentially as described by Kemp et al. (1977). Briefly, the reaction mixture (total volume 70 μl) contained the respective peptide at concentrations ranging from 0.1 to 1 mM in the following solution: 10 μCi [γ‐32P]ATP (0.5 mM), 62.5 mM MES, pH 6.9, 12.5 mM magnesium acetate, 0.25 mM EGTA and either protein kinase A (catalytic subunit, Boehringer, Mannheim, Germany) or muscle extracts prepared as described below. After incubation at 37°C for 0, 1, 2, 5, 10 and 15 min, 8 μl aliquots were removed from the reaction and the phosphorylated peptides were separated from [γ‐32P]ATP by the phosphocellulose binding technique described by Casnellie (1991). Apparent Km and Vmax values were determined by fitting the data of a double reciprocal Lineweaver–Burk plot to the Michaelis–Menten equation using the method of least squares.
Phosphoamino acid analysis
32P‐Labelled protein was recovered from sample buffer by the method of Wessel and Flügge (1984), dried and hydrolysed in 6 M HCl at 110°C for 2 h. After lyophilization, the hydrolysate was dissolved in 10 μl of H2O and applied to a Polygram CEL400 Uni layer plate (Merck, Darmstadt, Germany). Electrophoretic separation of phosphoamino acids was in 10% acetic acid, 1% pyridin, pH 3.5, at 800 V for 1 h. Radioactively labelled phosphoamino acids were identified by visualization of standard phosphoamino acids with ninhydrin and autoradiography.
Preparation of tissue extracts
Bovine or rat skeletal muscle, dissected into pieces, was shock‐frozen in liquid nitrogen and stored at −80°C. Then 0.5 g of material were homogenized in 3 vol of ice‐cold 5 mM EDTA, 15 mM 2–mercaptoethanol pH 7.0 and the resulting suspension clarified by centrifugation (4°C, 11 000 g, 10 min). The supernatant was applied onto a MonoQ HR 5/5 column (Pharmacia, Uppsala, Sweden). After washing with a buffer containing 50 mM Tris–HCl pH 7.9 and 1 mM DTT, protein was eluted with a linear gradient from 0 to 500 mM KCl in the same buffer. Assays for kinase activity were performed as described above.
Myosin binding assays
α‐Chymotryptic myosin subfragments (myosin rod and LMM) and recombinant LMM fragments (LMM 75, LMM 59, LMM50, LMM 50–75, LMM 30) were adjusted to 50 mM KCl, 5 mM Na phosphate pH 7.0 to allow the formation of filaments. Approximately 1 μg of each suspension was spotted onto nitrocellulose membranes (BA‐85, Schleicher and Schüll, Dassel, Germany). After air drying, the strips were blocked with overlay buffer (1% bovine serum albumin, 0.2% Tween‐20, 100 mM KCl, 20 mM imidazole‐HCl pH 7.0, 1 mM DTT) for 30 min. Individual strips were treated for 60 min with the respective myomesin fragments in the same buffer. After three washes with overlay buffer, strips were incubated for 45 min with rat monoclonal antibody YL1/2 (Wehland et al., 1984), specific for the EEF tag of our recombinant myomesin fragments. Strips were washed as described above and treated with peroxidase‐conjugated goat anti‐rat antibody (Dianova) diluted 1:1000 in overlay buffer for 30 min. After a final washing cycle, antibody binding was visualized by reaction in 5 ml of 100 mM Tris–HCl pH 7.5 supplemented with 100 μl of diaminobenzidine (40 mg/ml stock), 25 μl of NiCl2 (80 mg/ml stock) and 1.5 μl of 30% H2O2. All steps were performed at room temperature.
Titin binding assay
Recombinantly expressed titin fragments (0.2 μg for each domain) were spotted on nitrocellulose membranes (BA‐85, Schleicher and Schüll, Dassel, Germany) pre‐wetted with phosphate‐buffered saline. Blocking, incubation with T7‐tagged myomesin fragments and washing was as described for the myosin binding assays (see above). Binding to titin was detected by the T7 tag‐specific murine monoclonal antibody (1:1000 diluted, Novagen) followed by peroxidase‐conjugated goat anti‐mouse antibody (1:1000 diluted, Dianova) in overlay buffer. Visualization of antibody binding was as described for the myosin binding assay.
SDS–PAGE and immunoblotting were as described (Fürst et al., 1988). Protein concentrations were obtained with the BioRad dye reagent. Synthetic peptides were obtained by Fmoc chemistry on a Pep Synthesizer TM9050 (Millipore Co., Bedford, USA). HPLC analysis of synthetic peptides was performed on a Vydac 218 TP54 column at 50°C using a gradient of 0–90% acetonitrile in 0.1% trifluoroacetic acid as eluent.
Dr Alfred Wittinghofer (Max‐Planck‐Institute for Molecular Physiology, Dortmund, Germany) kindly provided the cDNA of rabbit myosin heavy chain 2/10. We thank Frank Steiner (this institute) for making the recombinant myosin constructs and Uwe Plessmann for expert technical assistance. This work was supported in part by grants from the Deutsche Forschungsgemeinschaft to D.O.F., M.G. and K.W.
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