Functional phosphorylation sites in cardiac myofilament proteins are evolutionarily conserved in skeletal myofilament proteins

Abstract

Protein phosphorylation plays an important role in regulating cardiac contractile function, but phosphorylation is not thought to play a regulatory role in skeletal muscle. To examine how myofilament phosphorylation arose in the human heart, we analyzed the amino acid sequences of 25 cardiac phosphorylation sites in animals ranging from fruit flies to humans. These analyses indicated that of the 25 human phosphorylation sites examined, 11 have been conserved across vertebrates and four have been sporadically present in vertebrates. Furthermore, all 11 of the cardiac sites found across vertebrates were present in skeletal muscle isoforms, along with three sites that were sporadically present. Based on the conservation of amino acid sequences between cardiac and skeletal contractile proteins, we tested for phosphorylation in mammalian skeletal muscle using several biochemical techniques and found evidence that multiple myofilament proteins were phosphorylated. Several of these phosphorylation sites were validated using mass spectrometry, including one site that is present in slow- and fast-twitch troponin I (TnI), but was lost in cardiac TnI. Thus, several myofilament phosphorylation sites present in the human heart likely arose in invertebrate muscle, have been evolutionarily conserved in skeletal muscle, and potentially have functional effects in both skeletal and cardiac muscle.

myofilament protein evolution has deep roots. Using comparative analysis from genomes of species representing metazoans, closely related protists, fungi, and other eukaryotic groups, Steinmetz et al. (27) reconstructed key steps in muscle evolution. They identified a core set of contractile proteins [actin, myosin heavy chain II (MHC), regulatory and essential myosin light chains, regulatory myosin light chain (RLC), essential myosin light chain (LC1), and tropomyosin (Tm)] that predates the evolution of muscle tissue. They further found that the troponin complex [troponin I (TnI), cardiac troponin (TnC), and troponin T (TnT)] is conserved among bilaterians. More recent genome duplications that occurred coincidentally with the evolution of vertebrate animals led to the emergence of multiple isoforms for different myofilament proteins (12, 17). This enabled vertebrate animals to express a different protein isoform in cardiac, slow- or fast-twitch muscle [TnI, TnT, myosin binding protein C (MyBPC)], or express one isoform in fast-twitch muscle and a second isoform in both ventricular and slow-twitch muscle (TnC, RLC, LC1, MHC).

Phosphorylation of the myosin RLC appears to be an ancient mode of muscle contraction regulation (27). Myosin light chain kinase first appeared in metazoan animals and allowed for the tight regulation of actomyosin contraction by coupling RLC phosphorylation to elevated cytoplasmic Ca²⁺ concentrations in muscle and nonmuscle cells (27). In human cardiac muscle, phosphorylation as a mode of regulation appears to have expanded, as there are at least 20 known serine and threonine residues in seven myofilament phosphoproteins whose phosphorylation/dephosphorylation have known functional consequences (9, 19, 29). Shaffer and Gillis (24) found an increased number of phosphorylation sites in cardiac isoforms of TnI and MyBPC in endothermic vertebrates, suggesting that as the heart became more complex so did the number of sites for regulation. Far fewer phosphorylation sites are known among the skeletal muscle isoforms of the same myofilament proteins (14, 18, 21, 30, 32), and their correspondence to cardiac sites has not been analyzed. The common ancestry of skeletal and cardiac myofilament proteins along with recent evidence of phosphorylation in skeletal myofilament proteins (6, 7, 13, 21) suggests a regulatory mechanism that is common between muscle types.

In the heart, contractility can be either increased or decreased by altering the phosphorylation pattern of different myofilament proteins (26). If the cardiac phosphorylation sites that alter contractile function arose in invertebrates and were then conserved in skeletal muscle, phosphorylation as a regulatory mechanism in cardiac muscle may also be present in skeletal muscle. Recently, we and others have shown that the posttranslational modification of cysteines in myofilament proteins can alter contractile function by increasing calcium sensitivity in fast twitch muscle and by increasing basal contractile activity in cardiac and slow-twitch muscle (5, 15). Thus, it is likely that posttranslational modifications to other amino acids could alter skeletal muscle contractile function.

In this investigation, we focused on 25 phosphorylation sites in five human cardiac myofilament proteins whose phosphorylation/dephosphorylation have known functional consequences. We first compared the amino acid sequences of these cardiac myofilament proteins to the corresponding sequences of homologs from animals that represent key branch points in muscle evolution, asking where in the phylogeny of animals the serine and threonine residues appeared. We found that the phosphorylation sites arose at all levels of the phylogenetic tree, from arthropods to mammals. We then asked whether the skeletal muscle paralogs of the cardiac phosphoproteins were potential analogous phosphoproteins. Fourteen of the 25 cardiac myofilament phosphorylation sites had serines or threonines in the corresponding skeletal myofilament sequence. Using one- (1D) and two-dimensional (2D) gel electrophoresis, we showed that many skeletal muscle paralogs were phosphorylated, and many had multiple charge variants. Finally, we used mass spectrometry to confirm phosphorylation sites in skeletal myofilament proteins and identified 11 sites, including one site that is present in slow- and fast-twitch TnI but was lost in cardiac TnI. Together with the discovery of many skeletal muscle phosphorylation sites and new insights into muscle and phosphorylation evolution, our data support a regulatory role of phosphorylation in skeletal muscle contractile function.

MATERIALS AND METHODS

Protein sequences and structures.

Amino acid sequences were collected from Uniprot, GenBank, and Ensembl databases and aligned using Clustal Omega (25). Percent sequence identity was determined using Clustal Omega alignments. Myofilament protein structures for human cardiac troponin (1J1E) and chicken fast troponin (1YVO and 1YTZ) were accessed from the PDB database and modeled using PyMol. Unrooted phylogenetic trees were constructed using MUSCLE to align sequences, Gblocks to eliminate poorly conserved regions, PhyML to create the phylogeny, and TreeDyn to create each tree (2). Cardiac phosphorylation sites were chosen for phylogenetic analysis based on their identification and reported ability to alter cardiac contractile function as defined and reviewed in references (9, 19, 29, 38).

Muscle and myofibrils preparations.

Striated muscle tissues were collected from New Zealand White rabbits and Long-Evans rats. Animals were donated by investigators whose animal care and use protocols were approved by the University of California Berkeley Animal Care and Use Committee. Immediately following dissection the extensor digitorum longus (EDL), soleus, psoas, and ventricles were minced and homogenized in rigor buffer. The rigor buffer was composed of 120 mM potassium acetate, 50 mM HEPES, 4 mM magnesium chloride, and 5 mM EGTA with a pH of 7.0. Following homogenization samples were centrifuged at 16.1 relative centrifugal force for 5 min, and the supernatant containing mostly cytosolic proteins was discarded. The pellet was then resuspended in rigor buffer containing 1% Triton X-100 to solubilize membranous proteins. The homogenate was again centrifuged, and the supernatant discarded. This step was repeated twice to produce a pellet highly enriched in myofilament proteins. Following the last centrifugation, the myofibril pellet was resuspended in a 50:50 mixture of rigor buffer and glycerol and stored at −20°C for future use. To alter protein phosphorylation myofibrils were resuspended in NEBuffer 3 [100 mM NaCl, 50 mM Tris·HCl, 10 mM MgCl₂, 1 mM dithiothreitol (DTT), pH 7.9 at 25°C] containing 1 unit/μl calf intestinal phosphatase (New England BioLabs) and incubated for 60 min at 37°C. Control samples were resuspended in buffer without added phosphatase. To assess calcium-dependent phosphorylation, single rat soleus fibers were isolated and mechanically skinned in paraffin oil and then incubated in rigor buffer containing 5 mmol/l ATP with or without added calcium (28).

Protein gel electrophoresis.

Myofilament proteins were separated by size and charge using gel electrophoresis. To separate proteins by charge we used isoelectric focusing (IEF) and nonequilibrium gel electrophoresis (NEPHGE). For analysis of proteins with acidic isoelectric points including the myosin light chains and Tm, we performed IEF in tube gels containing 5% acrylamide using ampholines pH 4–6.5. For proteins with basic isoelectric points including TnT and TnI, we used NEPHGE with a combination of ampholines pH 3–10 and 7–9. After focusing in the first dimension, we placed the tube gels in DTT for 15 min followed by iodoacetamide for an additional 15 min to reduce and block cysteine residues. The tube gels were then positioned at the top of 12.5% acrylamide SDS PAGE gels for separation of proteins based on their relative molecular weight. The 1D SDS-PAGE gels were composed of a 5% acrylamide stacking gel and a 12.5% resolving gel. For 2D gels 100 μg of protein were loaded, and for 1D SDS PAGE gels 15 μg of myofibril protein were loaded.

Following electrophoresis gels were placed in a fixing solution containing 50% methanol and 10% acetic acid for a minimum of 30 min and then repeatedly washed in water. Gels were stained overnight with the fluorescent protein stain Sypro Ruby (Life Technologies) and destained in solution containing 10% methanol and 7% acetic acid. Gels were scanned using a Typhoon fluorescent scanner, and proteins visualized using excitation and emission wavelengths of 488 nm and 610 nm respectively. For analysis of protein phosphorylation gels were stained with Diamond Pro-Q (Life Technologies) following the manufacturer's instructions and imaged using the Typhoon scanner at 532 nm excitation and 580 nm emission wavelengths. Gels were then stained with Sypro Ruby to image total protein levels. For immunoblot experiments, proteins were transferred from SDS-PAGE gels to PVDF membranes and blocked in Tris-buffered saline with 5% milk. Primary antibodies for TnI (TI-4) and TnT (CT3) were purchased from the DHSB and used at 1:100 dilution. An antibody for fast TnI (Clone JLT-12) was purchased from Sigma. To visualize proteins, membranes were incubated with horseradish peroxidase conjugated secondary antibodies followed by incubation with ECL solution (Amersham Bio).

Peptide collection for mass spectrometry.

Mouse C2 myoblasts (39) were grown for 24 h in DMEM containing 10% fetal bovine serum and 10% fetal calf serum. The medium was then replaced with DMEM containing 2% horse serum to induce differentiation. After 72 h whole protein lysates were collected in 8 M urea, 75 mM NaCl, 50 mM Tris, pH 8.2, and phosphatase inhibitors following the protocol of Villén and Gygi (36). Whole cell protein lysates were sonicated three times for 60 s at 4° C with 2 min incubation on ice in between. Samples were then treated with 5 mM DTT for 25 min at 55° C, 14 mM iodoacetamide for 30 min at 25° C, and 5 mM DTT for 15 min at 25° C. Next, the samples were either diluted with 25 mM Tris·HCl, pH 8.2, and supplemented with 1 mM CaCl₂ and trypsin or digested in the lysis buffer with Lys-C at 37°C. After 18 h digestion was stopped with trifluoroacetic acid (TFA) 0.4% (vol/vol). Samples were then centrifuged at 2,500 g for 10 min at room temperature, and the pellet was discarded. Peptides were desalted using reverse-phase C18 SepPak cartridges from Waters following the manufacturer's instructions and dried down using vacuum centrifugation. Phosphopeptides were enriched using the protocol from Ficarro et al. (3). Briefly, peptides were resuspended in 80% acetonitrile, 0.1% TFA, and activated magnetic Ni-NTA-agarose beads (Qiagen Valencia, CA) for 30 min with end-over-end rotation. The beads were then washed three times with 80% acetonitrile and 0.1% TFA. Phosphopeptides were then eluted using 1:1 acetonitrile/1:20 ammonia/water. The enriched phosphopeptides were then acidified with 10% TFA and dried down using vacuum centrifugation.

Liquid chromatography-mass spectrometry analysis of affinity purified phosphopeptides.

Peptides were dissolved in 22 μl of 5% formic acid and 20 μl injected onto an Acclaim PepMap 100 μm × 2 cm NanoViper C18, 5 μm trap on a switching valve. After 5 min of loading at 5 μl/min, the trap column was switched on-line to a PepMap RSLC C18, 2 μm, 75 μm × 25 cm EasySpray column (Thermo Scientific). Peptides were then separated using a Dionex NCS-3500RS UltiMate RSLCnano UPLC system (Thermo Scientific) using a 7.5–30% acetonitrile gradient in an aqueous mobile phase containing 0.1% formic acid at a 300 nl/min flow rate over 90 min. Tandem mass spectrometry data was collected using an Orbitrap Fusion Tribrid instrument configured with an EasySpray NanoSource (Thermo Scientific). Survey scans were performed in the Orbitrap mass analyzer, and data-dependent MS2 scans in the linear ion trap using a combination of higher energy collision-induced dissociation (HCD) and electron transfer dissociation (ETD) using a decision tree based on peptide charge state and m/z range.

Informatics analysis.

Results from trypsin and LysC digests were processed using Protein Discoverer 1.4 (Thermo Scientific) to identify phosphopeptides. MS2 spectra were searched using Sequest HT with a Swiss-Prot Mouse database downloaded Nov 2014 (Swiss Institute of Bioinformatics, Geneva, Switzerland) with a maximum of t missed cleavages, minimum peptide length of 6, max Delta Cn of 0.05, 10 ppm, and 1 Da mass tolerances for precursor and fragment ions, respectively, monoistopic mass calculation, dynamic modifications for phosphate (+79.966 Da for S, T, and Y) and oxidation (+15.995 Da for M), and static modification for carbamidomethylation (+57.021 Da for C). HCD data were searched by specifying y and b ion fragments, and ETD data by specifying c and z ion fragments. ETD MS2 spectra were processed using a nonfragment filter to remove precursor peaks and their charged reduced forms prior to Sequest HT searches. Peptide identifications were filtered with Percolator (10) using matches to a decoy database, and peptides with q scores > 0.01 removed, resulting in a peptide false discovery rate of < 1%. HCD and ETD data were independently analyzed using phosphoRS 3.0 to assign probabilities of phosphate localization to S, T, and Y residues in each peptide (34).

RESULTS

Most functional human cardiac myofilament phosphorylation sites arose in invertebrates.

To understand the evolution of phosphorylation sites in cardiac myofilament proteins, we compared 25 human cardiac phosphorylation sites (9, 19, 29) to corresponding sites from animals representative of different branches of a phylogenetic tree (1, 12, 17). The comparison revealed that of the 25 phosphorylation sites in human cardiac proteins six sites were unique to mammals (Fig. 1A). We found that 11 phosphorylation sites were present in cardiac myofilament proteins of all bony vertebrate representatives (mammals + birds + amphibians + teleost fish). Four phosphorylation sites, TnI Ser 166, TnI Thr 181, TnT Ser 208, and LC1 Ser 200, were sporadically present across vertebrate evolution (Fig. 1, B–E).

Fig. 1.

Multiple cardiac phosphorylation sites have been conserved across animal evolution. A: comparison of serine and threonine phosphorylation sites found in human cardiac myofilament isoforms across animal evolution. Red boxes indicate conservation of a serine or threonine residue in the majority of analyzed genomes for each respective grouping. Empty boxes indicate the lack of a serine or threonine in the majority of analyzed genomes. Sequence numbering is based on the respective human protein. B: aligned segment of cardiac troponin I (cTnI) sequences from an evolutionarily diverse range of vertebrate species. Ser 166 is highlighted in red. C: aligned cTnI sequences from a similar range of vertebrate species. Thr 181 is highlighted in red. D: aligned cardiac troponin T (TnT) sequences from a similar range of vertebrate species. Thr 204 and Ser 208 are highlighted in red. E: aligned cardiac essential light chain (LC1) sequences from a similar range of vertebrate species. Ser 200 is highlighted in red. TM, tropomyosin; RLC, regulatory myosin light chain.

All cardiac phosphorylation sites that arose in invertebrates are found in skeletal paralogs.

To further understand the evolution of phosphorylation sites, we asked if skeletal muscle paralogs of cardiac myofilament proteins had a corresponding serine or threonine. We first constructed phylogenetic trees for each protein and confirmed the common evolutionary origin and groupings between the different isoforms (Figs. 2, A and B, and 3, A and B). Then analyzing the sequences, we found that all of the sites that were present before the evolution of fiber types were also present in either slow- or fast-twitch isoforms, while none of the four cardiac sites that were absent in tetrapod predecessors (TnI Ser 23, TnI Ser 24, TnI Tyr 26, TnT Thr 204) was found in a skeletal muscle paralog (Figs. 2, 3, and 4). Nor were the six cardiac sites unique to mammals (TnI Ser 5, TnI Ser 6, TnI Ser 77, TnI Thr 78, TnI Thr 143, and TnT Thr 294). Note that cTnI Thr 143 is found in zebrafish but is absent in the majority of fish species analyzed (Fig. 4). Three of the four cardiac sites that were sporadically present across cardiac vertebrate evolution (TnI Ser 166, TnI Thr 181, LC1 Ser 200) were found in a skeletal muscle paralog. In summary, 14 of the 25 cardiac phosphorylation sites were present in sequences of a skeletal muscle paralog. The 11 cardiac sites that were not found in a skeletal muscle paralog are those that first appeared in amphibians or mammals or that were sporadically present across evolution.

Fig. 2.

Troponin T and troponin I phosphorylation sites are partially conserved across striated muscle evolution. A and B: unrooted phylogenetic tree of fast, slow, cardiac, and ancestral isoforms of troponin T and troponin I, respectively. Colored backgrounds were added to show the relationships between fiber type isoforms. C and D: sequence alignments of cardiac and fast- and slow-twitch isoforms from 4 evolutionarily diverse vertebrate species and 3 invertebrate species. Phosphorylation sites are listed above the respective sequences and numbered based on the respective human cardiac protein. Conserved serines or threonines are highlighted red. In TnT Ser 285, homology was poor for lancelet and fruit fly sequences, and we were unable to produce an accurate alignment. In the fast-twitch isoforms of TnI from birds and mammals there is a single threonine rather than the dual Ser 42 and Ser 44 sites in the cardiac and slow-twitch isoforms. Previous experiments have shown phosphorylation of this threonine in rabbit fast-twitch muscle (8, 14), and therefore we have highlighted this site. Percent sequence identity for each protein is based on comparison to the human cardiac protein and is shown on the far right.

Fig. 3.

Essential and regulatory light chain phosphorylation sites are partially conserved across striated muscle evolution. A and B: unrooted phylogenetic tree of fast, cardiac, atrial, and ancestral isoforms of the essential and regulatory myosin light chains, respectively. Colored backgrounds were added to show the relationships between fiber type isoforms. C: sequence alignments of ventricular, atrial, and fast isoforms of 4 evolutionarily diverse vertebrate and 3 invertebrate species. Ventricular isoforms are expressed in both ventricular and slow-twitch muscle. Phosphorylation sites (numbering based on the human cardiac protein) are listed above the respective sequences. Conserved serines or threonines are highlighted red. Serine 200 of LC1 is adjacent to the COOH-terminal amino acid of the protein. We were unable to locate an annotated sequence for the chicken atrial regulatory light chain. Percent sequence identity for each protein is based on comparison to the human cardiac protein and is shown on the far right.

Fig. 4.

Unique aspects of skeletal and cardiac myofilament sequence evolution. A: diagram of the NH₂-terminal intron and exon structure of TnI. Exons are drawn as rectangles and introns as black lines. At the NH₂ terminus of TnI the cardiac gene contains an exon (marked in blue) not present in the slow- or fast-twitch isoforms. Present in this exon are Ser 23, Ser 24, and Tyr 26, which are each conserved in amphibians, birds/reptiles, and mammals. B: cardiac Thr 143 from TnI is present in zebrafish but not in most other fish. C: sequence conservation of Ser 283 from Tm in vertebrates.

To further analyze phosphorylation sites in myofilament protein paralogs, we compared the crystal structures of human cardiac troponin with chicken fast-twitch troponin (33, 37). Superimposing the two structures shows that the relative locations and potential accessibility to kinases for the conserved troponin sites are similar between each paralog, although some sites differ in the trimeric complex due to differences between cardiac and skeletal isoforms of TnC (Fig. 5).

Fig. 5.

Locations of cardiac and skeletal phosphorylation sites in troponin. Crystal structure of human cardiac troponin (PDB: 1J1E) containing TnI (red), TnT (green), and TnC (blue) superimposed with the crystal structure from chicken fast troponin (PDB: 1YTZ) shown in gray. The locations of the conserved phosphorylation sites are labeled and colored black. Phosphorylation site numbering is based on the human protein sequences. cTnI Ser 44 equates to fTnI Thr 12, cTnI Thr 51 pairs with fTnI Ser 20, cTnI Ser 150 pairs with fTnI Ser 118, and cTnT Thr 213 pairs with fTnT Thr 181. fTnT Ser 253 pairs with cTnT Ser 285, although this site is absent in the cardiac structure.

Myofilament proteins are phosphorylated in skeletal and cardiac muscle.

The sequence and structural conservation between cardiac phosphorylation sites suggested that the skeletal myofilament protein paralogs were phosphorylated. Using a gel phosphorylation stain (Diamond Pro-Q) that is frequently used to asses phosphorylation in cardiac myofilament proteins (35, 40, 41) and a total protein stain (Sypro), we analyzed the phosphorylation status of muscle samples from rat ventricles, rabbit psoas, and rat soleus and EDL that were highly enriched with myofilament proteins (myofibril preparation). We found Pro-Q fluorescence, indicating phosphorylation of myofilament proteins, in fast and slow skeletal muscle samples, as well as ventricular samples (Fig. 6A). Rat ventricular muscle samples had Pro-Q fluorescence at molecular weights that indicated RLC, TnI, Tm, TnT, and MyBPC were phosphorylated. Rat skeletal muscle samples had Pro-Q fluorescence at molecular weights that suggested that the same myofilament proteins were phosphorylated, with the exception of TnI. Neither cardiac nor skeletal muscle samples showed Pro-Q fluorescence at a molecular weight expected for LC1. Note that the cardiac and skeletal isoforms for most myofilament proteins differ in molecular weight. For instance, the molecular weight of cardiac TnT is 34.5 kDa, whereas fast skeletal TnT has multiple splice variants, with molecular weights centered around 30 kDa.

Fig. 6.

Ventricular and skeletal myofilament proteins are phosphorylated. A: SDS-PAGE gel stained with Pro-Q to identify phosphorylated proteins in myofibril samples from rat ventricles, rat soleus, rat extensor digitorum longus (EDL), and rabbit psoas muscle (2 biological replicates for each muscle type). Arrows to the right illustrate the molecular weight. The same Pro-Q stained gel was subsequently stained with Sypro to visualize the total protein content. B: SDS-PAGE gels sequentially stained with Diamond Pro Q and Sypro Ruby showing that treatment of rat EDL myofibrils with alkaline phosphatase (AP) reduces the phosphorylation of TnT, TM, and an unidentified high-molecular-weight protein (highlighted by surrounding box). C: incubation of mechanically skinned rat soleus muscle fibers in a calcium rich solution increases the phosphorylation of RLC above levels in a low calcium solution (control) (highlighted by surrounding box) as visualized using SDS-PAGE gels with Pro-Q phosphorylation staining followed by Sypro total protein staining.

Phosphorylated proteins should have a change in Pro-Q fluorescence when treated with a kinase or phosphatase. We therefore treated skeletal myofibrils with alkaline phosphatase and found the fluorescence of the three bands for Tm and TnT were decreased 5.1-, 5.2-, and 7.8-fold (n = 5) compared with control myofibrils confirming these skeletal muscle proteins were phosphorylated (Fig. 6B). We next tested for the calcium dependence of phosphorylation, by mechanically skinning single fibers in oil and then bathing the fibers in a calcium solution. This increased the fluorescence of the RLC band 12.7-fold (n = 3) compared with the band from control fibers (Fig. 6C). We did not observe significant fluorescence changes in other myofilament proteins. In summary, comparison of samples with and without added alkaline phosphatase revealed changes in phosphorylation of TnT, Tm, and a high-molecular-weight protein, and comparison between samples in low and high Ca²⁺ confirmed calcium-dependent phosphorylation of skeletal RLC.

The Diamond ProQ staining suggested that myofilament proteins in both cardiac and skeletal myofibrils were phosphorylated (Fig. 6A). To further analyze phosphorylation in skeletal muscle, we used 2D electrophoresis, which separates proteins first by charge and then by molecular weight. Phosphorylation adds a negative charge, making 2D gels effective for identifying phosphorylated proteins, although other posttranslational modifications can also alter protein charge. Additionally, 2D gels provide an indication of the relative stoichiometry of posttranslational modifications and also allow separation of comigrating proteins present in 1D gels. Using pH 4–6.5 IEF gels, we identified charge variants suggestive of phosphorylation in the myofilament proteins with acidic isoelectric points: Tm, RLC, and LC1 (Fig. 7A). To separate TnT and TnI charge variants, we used pH 3–10 NEPHGE gels (Fig. 7A). TnT is alternatively spliced and therefore has protein variants that vary in both isoelectric point and molecular weight. To identify TnT and confirm its dephosphorylation, we performed immunoblots on control fast skeletal myofibrils and myofibrils treated with alkaline phosphatase (Fig. 7, B and C). The immunoblots revealed multiple charge variants and the expected shift in the protein spot pattern after phosphatase treatment. We also confirmed two fTnI charge variants using immunoblots (Fig. 7C). In summary, 2D gels confirmed charge variants consistent with phosphorylation for skeletal RLC, TnT, and Tm and further suggested phosphorylation of fast skeletal LC1 and TnI.

Fig. 7.

2D gels reveal multiple charge variants in myofilament proteins. A: rat ventricular and rat EDL (Skeletal) myofibrils focused using NEPHGE (left column pH 3–10) and IEF (right column pH 4–6.5) followed by SDS-PAGE electrophoresis and visualized with Sypro Ruby fluorescent protein stain. Selected myofilament proteins and their charge variants are labeled. B: immunoblots of TnT from 2D gels. In the top blot rat EDL muscle was incubated in alkaline phosphatase (AP) buffer without phosphatase. In the bottom blot the muscle sample was incubated in AP buffer with alkaline phosphatase resulting in a reduction in the number and relative amount of acidic charge variants in TnT. C: immunoblot of TnT and TnI from rat EDL muscle revealing multiple TnI and TnT charge variants and confirming their identification in the 2D gels.

Mass spectrometry confirms skeletal myofilament phosphorylation and identifies several additional sites.

Since the phylogenetic and biochemical data suggested that skeletal myofilament proteins were phosphorylated with relatively high stoichiometry, we next attempted to validate the phosphorylation sites using mass spectrometry. Following phosphopeptide enrichment, we identified several of the skeletal phosphorylation sites predicted from their sequence conservation in RLC, Tm, and TnI (Fig. 8A). We also identified sites in atrial LC1 (Thr 62, equivalent to Thr 69 in Fig. 3). RLC Thr 114 and TnI Ser 57 phosphorylation sites have not been previously identified. The predicted skeletal phosphorylation sites that we did not find often had nearby arginines and lysines, making the peptides produced from trypsin and Lys-C digestion short and unlikely to be sampled by the mass spectrometer. In analyzing the sequence conservation of the phosphorylated peptides, we identified one site (Ser 57 in fTnI) that is absent in human cardiac muscle, but is present in ancestral, fast-, and slow-twitch isoforms (Fig. 8B). In the regulatory light chain, we also identified a second phosphorylation site adjacent to Ser 15 that is present in vertebrate fast isoforms and in sea squirts as a threonine but is absent in the cardiac/slow-twitch isoforms (Figs. 3C, 8A). The other phosphorylation sites identified (fRLC: Thr 114, sTnI: Ser 183) were not conserved across vertebrate evolution. In summary, using mass spectrometry we validated multiple phosphorylated peptides and identified several sites that were unique to skeletal muscle isoforms. These data provide evidence of the conservation, gain, and loss of phosphorylation sites across muscle types, suggesting a broad physiological mechanism for regulating contractile function (Fig. 9).

Fig. 8.

Identification of phospho-peptides in skeletal myofilament proteins. A: table listing the myofilament protein (gene name in parentheses) followed by the peptide sequence identified through mass spectrometry (phosphorylation site is in lowercase). Phosphorylation site numbering is based on the mouse protein sequence. The calculated phosphoRS probability score for each phosphorylation site (34), and references to figures showing the sequence conservation of each site are shown in the last 2 columns. All sites identified had a q-score < 0.01 (10). B. Sequence alignments for fTnI Ser 57 that includes cardiac and fast- and slow-twitch isoforms from 4 evolutionarily diverse vertebrate species and 3 invertebrate species. Phosphorylation sites identified using mass spectrometry and conserved serines are highlighted red.

Fig. 9.

Myofilament phylogeny. Phylogenetic tree illustrating the estimated point in animal evolution where serines or threonines that are phosphorylated in mammalian skeletal or cardiac muscle first appear. *Site identified in the mass spectrometry screen.

DISCUSSION

Protein phosphorylation is known to modify contractile function at many myofilament sites in the heart, but phosphorylation is far less known in skeletal muscle. In this investigation, we have found new evidence of phosphorylation as a potential regulatory mechanism in skeletal as well as cardiac myofilament proteins. Starting with a list of cardiac myofilament phosphorylation sites that modify cardiac contractile function, we have investigated corresponding skeletal muscle homologs. We found 14 skeletal sites with analogous cardiac functional phosphorylation sites and found evidence that 12 appeared before separate cardiac and skeletal muscle types evolved. Using both 1D and 2D SDS-PAGE gels and mass spectrometry, we found that skeletal isoforms of Tm, LC1, RLC, TnT, and TnI can be phosphorylated and also discovered new skeletal phosphorylation sites. Our data, with new findings about muscle evolution and phosphorylation site discovery, suggest a novel mechanism to alter skeletal muscle contractile function.

By tracing the phylogeny of 25 known functional phosphorylation sites in five cardiac myofilament proteins, we have found evidence suggesting that these sites arose throughout animal evolution, rather than evolving only among vertebrates in cardiac isoforms (Figs. 1, 9). By comparing amino acid sequences of skeletal homologs with sequences from the corresponding cardiac sequences, we have found that 14 of the 25 cardiac phosphorylation sites have corresponding phosphorylation sites in skeletal paralogs (Figs. 2–4). Sequence evidence indicated that these skeletal phosphorylation sites evolved early on. Of the 14 sites, 12 were present in representative invertebrate animals; the other two were present in vertebrate representatives from zebrafish on. The 11 sites without corresponding skeletal phosphorylation sites arose later than those with skeletal muscle paralogs or were sporadically present. This result is consistent with those of Shaffer and Gillis (24), who found an increased number of phosphorylatable sites in cTnI and cMyBPC in endothermic vertebrates in their phylogenetic analysis of TnI and MyBPC amino acid sequences.

Using protein phosphorylation stains, 2D gels, and mass spectrometry, we have demonstrated that the skeletal isoforms of Tm, LC1, RLC, TnI, and TnT can be phosphorylated. (Figs. 6–8). Gel phosphorylation stains and 2D gels (Figs. 6, 7) suggested that Tm, RLC, TnT, and MyBPC from fast and slow rat skeletal muscles were phosphorylated, and 2D gels (Fig. 7) further suggested that LC1 and TnI from fast rat skeletal muscles were phosphorylated. Addition of alkaline phosphatase decreased phosphorylation of Tm and TnT, and addition of Ca²⁺ increased phosphorylation of RLC (Fig. 6). Thus, each of our core myofilament proteins from fast skeletal muscle can be phosphorylated. Our mass spectrometry results validated several phosphorylation sites we had inferred from amino acid sequences and revealed additional skeletal phosphorylation sites not previously annotated. (Fig. 8, fRLC Thr 114, fTnI Ser 57). The sites, common to the Fig. 1 cardiac sites and the Fig. 8 mass spectrometry sites, are distributed in the phylogenic tree like the other skeletal phosphorylation sites (Fig. 9). Approximately half are present in invertebrate representatives, and the other half in vertebrate representatives.

Recently, others have also used advances in mass spectrometry to discover phosphorylation sites in skeletal myofilament proteins. Hoffman et al. (6) performed a global phosphoproteomic analysis of human skeletal muscle and L6 myotubes in response to exercise or 5'-AMP-activated protein kinase (AMPK) stimulation. They identified several myofilament phosphorylation sites analyzed here including four sites whose level of phosphorylation increased after exercise (sTnI Ser 135 and S169 equivalent to cTnI Ser 166 and Ser 199, and also fTnT Ser 166 and Ser 167: two sites that are unique to the fast isoform). This follows on the work of Sancho Solis et al. (21), who used top-down high-resolution mass spectrometry to demonstrate that isoforms of both cardiac and fast skeletal TnI proteins are readily phosphorylated by AMPK at cTnI Ser 150 and fTnI Ser 118. These studies are in line with a growing literature of phosphorylation site identification in cardiac and skeletal muscle (6, 13), and also the quantification of changes in protein phosphorylation from different states including heart failure (22, 23, 42) and diabetes (43). Thus, there is growing evidence of extensive phosphorylation of myofilament proteins and changes to their phosphorylation status in response to different physiological conditions.

Cardiac muscle uses myofilament phosphorylation to alter contractile function in a rapid and specific manner, but skeletal muscle function is thought to only be modified by central nervous system input and metabolites (4). The effects from phosphorylation in cardiac muscle are dependent on the kinases activated and the resultant sites phosphorylated. In broad terms, PKC tends to phosphorylate sites that reduce contractile function (29, 31), PKA sites tend to increase power (11, 26), and AMPK sites tend to increase the contractile efficiency (16). Our analysis indicates that at least 14 cardiac phosphorylation sites are conserved in skeletal isoforms, and based on their effect in cardiac muscle these sites have the potential to cause muscle fatigue (i.e., PKC sites cTnT Thr 213 and Ser 285) or potentiation (i.e., AMPK TnI sites discussed above). Thus, our investigation strongly suggests a series of mechanisms to increase or decrease skeletal muscle function depending on the situation and kinases activated.

Our phylogenetic analysis indicates that phosphorylation of cardiac myofilament proteins was not a trait that solely arose in vertebrates. Rather, myofilament phosphorylation likely arose in invertebrate muscle and has been conserved in both skeletal and cardiac muscle. If phosphorylation did arise in invertebrate muscle, we speculate that phosphorylation provided a mechanism independent of the central nervous system to modulate contractile function. In this manner, contractile efficiency could be modulated as occurs in the mammalian heart when Ser 150 of TnI is phosphorylated (16), or contractile strength could be altered as occurs after beta-adrenergic stimulation in the heart (11). These modifications could provide a selective advantage by allowing muscle function to be maintained or modulated in times of stress or under different environmental conditions.

The conservation of serines and threonines in mammals does not confirm that these sites are phosphorylated in all animal species or that contractile function is altered upon phosphorylation. Kinases require specific structural environments in their protein substrates for binding and phosphorylation. Thus, the critical characteristic may not be the evolution of a phosphorylation site, but rather the evolution of a structural difference that enabled functional effects from phosphorylation. Additionally, some phosphorylation sites have no known effect on contractile function or are constitutively phosphorylated. For example, three cardiac TnT phosphorylation sites (Thr 204, Ser 208, Thr 294) do not alter function upon phosphorylation (31). The absence of functional effects may be one explanation for the poor conservation in these three TnT sites across vertebrates. From our studies it can also be seen that phosphorylation sites have not only been gained across vertebrate evolution, but also lost in different muscle isoforms (fTnI Ser 57) (Fig. 8). As a consequence, it is likely that different animal species may have their own unique phosphorylation sites and that sites conserved in other vertebrates such as fish and reptiles have been lost in the mammalian lineage.

Phosphorylation is a well-studied mechanism for modulation of contractile function in cardiac muscle. Phosphorylation of skeletal muscle myofilament proteins and modulation of its contractile function is relatively unstudied. Recently, advances in mass spectrometry have enabled discovery of many phosphorylation sites in skeletal as well as cardiac myofilament proteins. In this investigation we sought evidence of skeletal muscle myofilament phosphorylation analogous to phosphorylation of cardiac myofilament proteins. We have shown that the cardiac phosphorylation sites arose throughout the course of animal evolution, not just after the whole genome duplications that occurred before vertebrate evolution. Of the 25 cardiac myofilament phosphorylation sites we studied, 14 could be found in the corresponding amino acid positions in the skeletal myofilament protein homologs. Finally, we used mass spectrometry to validate several of the phosphorylation sites inferred from amino acid sequences and found several additional skeletal phosphorylation sites. Together with the discovery of skeletal muscle phosphorylation sites and new insights into muscle and phosphorylation evolution, our data support a regulatory role of phosphorylation in skeletal muscle contractile function.

GRANTS

This work was funded by an Integrative Biology departmental summer fellowship. Mass spectrometric analysis was performed by the OHSU Proteomics Shared Resource with partial support from National Institutes of Health core grants P30EY-010572, P30CA-069533 and S10OD-012246.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

S.M.G. and S.L.L. conception and design of research; S.M.G. performed experiments; S.M.G. and S.L.L. analyzed data; S.M.G. and S.L.L. interpreted results of experiments; S.M.G. and S.L.L. prepared figures; S.M.G. and S.L.L. drafted manuscript; S.M.G. and S.L.L. edited and revised manuscript; S.M.G. and S.L.L. approved final version of manuscript.