Single Muscle Fiber Proteomics Reveals Distinct Protein Changes in

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Single muscle fiber proteomics reveals distinct protein changes in slow and fast fibers during muscle atrophy Franziska Lang, Solmaz Khaghani, Clara Tuerk, Janica Lea Wiederstein, Soraya Hölper, Tanja Piller, Leonardo Nogara, Bert Blaauw, Stefan Günther, Stefan Müller, Thomas Braun, and Marcus Krüger J. Proteome Res., Just Accepted Manuscript • DOI: 10.1021/acs.jproteome.8b00093 • Publication Date (Web): 24 Aug 2018 Downloaded from http://pubs.acs.org on August 26, 2018

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Journal of Proteome Research

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Single muscle fiber proteomics reveals distinct protein changes in slow and fast fibers during muscle atrophy Franziska Lang1,#, Solmaz Khaghani2,#, Clara Türk1, Janica Lea Wiederstein1, Soraya Hölper3, Tanja Piller4, Leonardo Nogara5, Bert Blaauw5, Stefan Günther2, Stefan Müller6, Thomas Braun2, Marcus Krüger1,

6,* #

contributed equally, * corresponding author

1

Institute for Genetics, Cologne Excellence Cluster on Cellular Stress Responses in Aging‐Associated Diseases

(CECAD), 2

Max Planck Institute for Heart and Lung Research, Ludwigstr. 43, 61231 Bad Nauheim, Germany

3

Sanofi-Aventis Deutschland GmbH, Biologics Research, Protein Therapeutics, Industriepark Höchst, 65926

Frankfurt, Germany 4

5

Institute of Biochemistry II, Goethe University Medical School, 60590 Frankfurt, Germany Venetian Institute of Molecular Medicine (VIMM), Department of Biomedical Sciences Padova, University of

Padova, Italy 6

Center for Molecular Medicine (CMMC), University of Cologne, 50931 Cologne, Germany

1 ACS Paragon Plus Environment


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ABSTRACT

Skeletal muscles are composed of heterogeneous collections of fibers with different metabolic profiles. With varied neuronal innervation and fiber type compositions, each muscle fulfils specific functions and responds differently to stimuli and perturbations. Here, we assessed individual fibers by mass spectrometry to dissect protein changes after loss of neuronal innervation due to section of the sciatic nerve in mice. This proteomics approach enabled us to quantify ~600 proteins per individual soleus and EDL (extensor digitorum longus) muscle fiber. Expression of myosin heavy chain (MyHC) in individual fibers enabled clustering of specific fiber types; comparison of fibers from control and denervated muscles with the same MyHC expression revealed restricted regulation of a total of 240 proteins in either type I, IIa or IIb fibers 7 days after denervation. The levels of several mitochondrial and proteasomal proteins were significantly altered, indicating rapid adaption of metabolic processes after denervation. Furthermore, we observed fiber type-specific regulation of proteins involved in calcium ion binding and transport, such as troponins, parvalbumin and ATP2A2, indicating marked remodeling of muscle contractility after denervation. This study provides novel insight into how different muscle fiber types remodel their proteomes during muscular atrophy.

KEY WORDS

Denervation, muscle atrophy, single muscle fiber, calcium signaling


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INTRODUCTION

Each skeletal muscle is composed of a heterogeneous collection of fiber types with distinct physiological adaption in response to a stimulus. For example, slow type I fibers contract slowly and are resistant to fatigue, whereas fast fibers contain a more efficient sarcoplasmic reticulum and contract faster (1). Each fiber has a specific molecular signature of myosin heavy chain molecules (MyHC). Slow type I fibers express mainly MYH7, while fast type fibers can be separated into three classes: type IIa (MYH2), IIx (MYH1) and IIb (MYH4) (2). Moreover, immunostaining and immunoblotting has revealed the existence of both pure and mixed fibers containing different combinations of MyHC molecules (3, 4); this knowledge has extended our interpretation of the physiological functions of muscle tissue. MyHCs are currently the best marker proteins for delineation of individual fiber types and several largescale transcriptome and proteome profiling studies have helped to dissect the molecular signature of whole muscles and individual fibers under regular and perturbed conditions (59). Another important feature of skeletal muscle tissue is its ability to respond rapidly to environmental changes. For example, physical exercise elevates protein synthesis (10, 11), whereas, starvation, disuse atrophy, aging or neurodegenerative diseases induce muscle wasting within a few days (12). Loss of neuronal stimulation results in reduced protein synthesis and enhanced protein breakdown, leading to a net loss of muscle tissue (13). However, the fast extensor digitorum longus muscle (EDL) exhibits increased protein synthesis at 7-10 days after denervation, which temporally diminishes induction of muscular atrophy (14). Although numerous studies on muscle denervation have been conducted, the exact physiological mechanisms by which single fibers with different MyHC protein expression and metabolic profiles alter their proteomes in response to denervation have been barely elucidated.



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Section of the sciatic nerve is an effective, established model system for studying cellular remodeling processes during muscle atrophy in rodents (15). The subsequent reduction in electrical signals and neurotrophic factors induces rapid remodeling of gene and protein expression (16, 17). Increased degradation of proteins and organelles is mainly mediated by activation of the ubiquitin-proteasome system. For example, different E3 ubiquitin ligases such as MurRF1 (Muscle-specific RING finger protein 1) and MAFbx/Atrogin-1 (Muscle atrophy F-box protein) are upregulated on both mRNA (18, 19) and protein level in muscle atrophy. Atrogin-1 is responsible for the degradation of transcription factors, whereas MuRF1 targets the myofibrillar proteins of thick filaments (20, 21). In addition, activation of the autophagy system is another important catabolic pathway that contributes to degradation of proteins under cellular stress and muscle atrophy (12). Earlier studies mainly assigned individual fiber types by immunohistochemistry using antibodies against MyHC isoforms, electrophoretic separation, succinate dehydrogenase staining, or myofibrillar ATPase activity (22-24). This techniques enables the classification of skeletal muscles as slow and fast muscles based on their type I and II fiber composition. For example, the gastrocnemius (GAST) and tibialis anterior (TA) muscles of the hind limb are comprised of a mixture of type I, IIa, IIb and IIx fibers, whereas the slow soleus muscle mainly contains an equal proportion of type I and type IIa fibers. The extensor digitorum longus (EDL), one of the fastest muscles in the mouse, is mainly comprised of type IIb fibers (2, 24, 25). To enable large-scale profiling of individual muscle fibers, recent studies have performed microgenomics on single fibers with different metabolic activities to identify regulatory genes that are preferentially expressed in specific fiber types (9). Modern high-sensitivity mass spectrometers can analyze samples containing low amounts of protein; single skeletal muscle fibers can even be assessed using shotgun liquid chromatography-mass spectrometry


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(LC-MS) (4, 7). Thus, mass spectrometric analysis of single muscle fibers represents a versatile approach that increases the resolution of skeletal muscle analyses. This could help to decipher the dynamics of fibers with different metabolic profiles under any pathological condition. Here, we employed a mouse model of section of the sciatic nerve to study denervation-induced muscle atrophy in individual fibers of hind limb muscles. Isolated muscle fibers were grouped based on MyHC protein expression, which allowed us to compare fiber type-specific response of soleus and EDL under normal and denervated conditions.



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EXPERIMENTAL SECTION Denervation and tissue lysis

Adult wild-type mice (C57BL/6J) were anaesthetized with an intraperitoneal injection of 100 mg/kg ketamine and 10 mg/kg xylazine. The denervation was performed by removing ~5 mm of sciatic nerve from the left hind limb. The right hind limb served as the control leg. Wounds were closed by suturing with 5-0 Vicryl. Mice were sacrificed 7 days after denervation and the soleus, extensor digitorum longus (EDL), tibialis anterior (TA) and gastrocnemius (GAST) muscles were isolated from the denervated and control hind limb. Isolated muscles were snap frozen in liquid nitrogen. Frozen tissue was cryogenically ground using a mortar and pestle and dissolved in cold lysis buffer (modified RIPA: 50 mM Tris/HCl pH 7.5, 150 mM NaCl, 1% NP-40, 1 mM EDTA, 0.1% sodium deoxycholate) with protease inhibitor cocktail. After homogenization on a rotating wheel for 30 min at 4 °C followed by sonication, protein extracts were centrifuged at 15,000 rpm for 10 min at 4 °C. Protein concentrations were determined using the PierceTM 660 nm Protein Assay (Thermo Scientific). All mouse experiments were performed in accordance with the regulations issued by the Committee for Animal Rights Protection of the State of Hessen, Germany.

Collagenase digestion and myofiber isolation

For proteomic analysis of single muscle fibers, denervated and control soleus and EDL muscles were isolated from euthanized mice and incubated with 0.2% collagenase P in DMEM media at 37 °C for approximately 20 to 40 min (depending on muscle size), as previously described (26). Each sample was assessed regularly to prevent over-digestion. The digestion was stopped by transferring the muscles into warm DMEM, then the muscles were carefully triturated with media using a Pasteur glass pipette to release myofibers. Fibers were transferred into a glass centrifuge tube, washed three times with PBS and single fibers were manually isolated under a stereomicrospcope (Leica). Proteins were extracted from single 6 ACS Paragon Plus Environment

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fibers by lysing the cells with 10 µL urea buffer (6 M urea, 2 M thiourea in 10 mM HEPES, pH 7.6). The muscle proteome reference dataset was generated by isolation of ~150 fibers from the diaphragm, EDL, flexor digitorum brevis (FDB), gluteus maximus (Glut), soleus, TA and GAST. After pooling ~150 muscle fibers, protein extraction and digestion were performed as described for the single fibers.

In-solution digestion

Protein digestion of muscle lysates, single fibers and pooled fibers was performed as previously described (27). Briefly, protein extracts were precipitated with ice-cold acetone at -20 °C for 2 h and resuspended in 10 µL of 8 M urea buffer. Then, samples were reduced with 10 mM dithiothreitol (DTT) for 30 min, followed by carbamidomethylation with 55 mM iodacetamide (IAA) in the dark for 30 min at room temperature. Proteins were digested with LysC for 2 h and then digested overnight with trypsin. The digestion was stopped by adding an equal amount of Buffer C (5% ACN, 1% trifluoroacetic acid) and the peptides were desalted using in-house made C18-based Stop and Go Extraction Tips (28).

Immunohistochemistry Immuno-histological stainings on muscle cryo-sections were performed as described previously (29). The following antibodies were used: ATP2A2/SERCA2 (Abcam, ab2861), CRYAB (Enzo, ADI-SPA-222), MYH7 (DSHB, BA-F8) and Alexa Fluor secondary antibodies (Invitrogen). Relative fluorescence intensities in individual fibers were determined using ImageJ by manually selecting the intracellular area of each fiber. Calculated intensity values were normalized to the controls.



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Immunoblotting Single muscle fibers were dissolved in 10 µL loading buffer (1x Laemmli buffer, 50 mM DTT) and heated for 10 min at 70°C. Fiber lysates were separated on TGX Acrylamide gels and transferred to polyvinylidene fluoride (PVDF) membranes by semi-dry blotting using the Bio-Rad Trans-Blot Turbo system. Fiber type was determined by using 1 µL of the lysate for SDS-PAGE analysis of MyHC expression and the remaining 9 µL of lysate were then used for fiber-type specific immunoblot analysis. Stain-Free loading controls and proteins levels visualized by chemiluminescence using ECL Reagent (Bio-Rad) were imaged with the ChemiDoc imaging system (Bio-Rad). The following antibodies were used: CRYAB (Enzo, ADI-SPA-222), MYH7 (DSHB, BA-F8; for fiber type identification) and anti-rabbit HRPcoupled secondary antibody (Sigma).

LC-MS/MS analysis

Mass spectrometric analysis was performed using an Easy nLC 1000 UHPLC coupled to a QExactive mass spectrometer (Thermo Fisher), as previously described (30). Peptides were fractionated using in-house made 50 cm columns packed with 1.9 µm C18 beads using a binary buffer system, consisting of Buffer A (0.1% FA) and Buffer B (80% ACN in 0.1% FA). All samples were analyzed over a 240 min gradient, raising the content of Buffer B from 10% to 38% over 210 min, then from 38% to 60% over 10 min, followed by washing with 95% Buffer B and re-equilibration with 5% Buffer B for 10 min.

MS data processing

The acquired raw data were analyzed using MaxQuant software and the implemented Andromeda software (1.5.2.8 and 1.5.3.8) (31). A mouse UniProt database (30.01.2015) with common contaminants was used for peptide and protein identification (32). All parameters in


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MaxQuant were set to the default values, trypsin was selected as the digestion enzyme and a maximum of two missed cleavages were allowed. Methionine oxidation and N-terminal acetylation were set as variable modifications and carbamidomethylation of cysteines was chosen as fixed modification. For analysis of MyHC isoforms, only unique peptides were used for quantification. A false discovery rate (FDR) of 1% was used for both the peptidespectrum matches and protein level matches with the implemented target decoy algorithm. All single fiber samples were analyzed together with seven raw files from the reference dataset and “the match between runs” option was enabled to quantify peptides with a missing MS/MS spectrum in the analysis of single soleus and EDL fiber samples. In addition, the option “separate LFQ parameter groups” was selected to distinguish between peptides from the single fiber analysis and peptide reference dataset.

MyHC fiber type assignment Fiber types were assigned based on the abundance of myosin (MyHC) isoforms as described in (7). The relative amount of each MyHC was determined by dividing the LFQ intensity of the respective isoform (MYH1, MYH2, MYH4, MYH7) by the sum of the intensities of all four MyHC isoforms. Fibers were classified as type I if the relative abundance of MYH7 was higher than 80%, as type IIa, if MYH2 > 60%; type IIx, if MYH1 > 60%; and type IIb, if MYH4 > 80% (7).

Data analysis Data analysis and visualization was performed using Perseus software and the R environment (33). Two-sided t-tests were used to identify differentially expressed proteins between groups defined by muscle fiber typing. t-test was applied to proteins with at least 3 LFQ intensity values per group. Multiple testing correction using a FDR of 1% and 5% was based on



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significance analysis of microarrays (SAM) (34). Here, 500 permutations and a fudge factor of 0.1 were chosen and a fold-change of 1.5 as a cut-off. Principal component analysis (PCA), gene ontology (GO) annotation and 2D-enrichment were carried out using Perseus software with the default settings (35).

RESULTS Distinct protein changes during denervation-induced atrophy in intact slow and fast skeletal muscles

First, we sought to determine the systematic effects of denervation-induced muscular atrophy at seven days after section of the sciatic nerve. We performed unilateral section of the sciatic nerve in C57BL/6J wild-type mice. We isolated several muscles including the soleus, EDL, GAST and TA from both the control and denervated hind limb. Three mice were assessed in each experiment (n = 3). After protein extraction, each sample was subjected to in-solution digestion and analyzed using a 4 h LC-MS gradient (Figure 1A).


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Figure 1: Skeletal muscles exhibit varied protein changes after neuronal denervation

(A) TA, GAST, soleus and EDL muscles from the control and denervated legs were isolated 7 days after denervation (upper panel). Proteins were digested with trypsin, subjected to a 4 h LC-MS/MS gradient and analyzed (lower panel). (B) Pearson correlation matrix of Euclidean distances between control and denervated muscles in biological triplicate (n = 3). The correlation is based on the log2 ratios of LFQ intensities. (C) 2-Dimensional enrichment of GO- and KEGG terms using the ratios in denervated and control (den/ctrl) soleus and EDL muscles. (D) Log2 den/ctrl ratio distribution of proteasomal proteins for different muscles. (E) Differentially regulated proteins in intact skeletal muscles were identified by plotting the -log10 P-values and log2 fold changes between control and denervated samples. Volcano plots with permutation based multiple testing correction at a FDR of 5% identified 319 (13%) differentially expressed proteins after denervation in the soleus, (F) 164 (9%) in the EDL, (G) 300 (15%) in the GAST and (H) 137 (9%) in the TA. (I) Venn diagram showing the low degree of overlap in the significantly regulated proteins between all four muscles.

We identified 50,581 unique peptides, resulting in 3,590 quantified proteins with a false discovery rate (FDR) below 1% at the protein and peptide level (Table S-1). Pearson correlation clustering based on Euclidian distances showed high reproducibility between biological triplicates and demonstrated that each muscle exhibited a different response to the loss of neuronal innervation (Figure 1B). To highlight differentially regulated pathways, we applied two-dimensional enrichment of the log2 protein fold changes between the denervated and control slow soleus and fast EDL muscles. We found that proteins involved in metabolic processes such as glycolysis and gluconeogenesis (KEGG pathway: hsa00010) were more significantly altered in the EDL than the soleus at one week after denervation (Figure 1C). Conversely, gene


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ontology (GO)-terms such as fatty acid metabolism and β-oxidation were markedly downregulated in the soleus. In addition, we observed several proteasomal subunits were expressed at enhanced levels in the EDL, TA and GAST at one week after denervation, but were not altered in the soleus (Figure 1D). To identify proteins that were differently regulated in control and denervated muscles, we performed a two-sided t-test using multiple testing correction based on significance analysis of microarrays (SAM) with a false discovery rate (FDR) of < 5% (34). 164 proteins were upregulated and 155 proteins were downregulated in the denervated soleus compared to the control soleus (Figure 1E). Significantly fewer proteins were downregulated in the EDL and TA after denervation (Figure 1F-H). Next, we compared the significantly differently regulated proteins in all four muscles, and identified 13 commonly regulated proteins (Figure 1I; Table S-1).

The number of exclusively regulated proteins ranged from 31 in the TA to 117 in the soleus (Figure 1I), indicating that each muscle - depending on its fiber content -exhibits a specific response to the loss of neuronal innervation (25, 36, 37).

Single fiber proteomics enables detection of MyHC isoforms and fiber type profiling

Next, we asked whether we could resolve the heterogenic response of different muscles to denervation by a more focused analysis of individual muscle fibers. Thus, at 7 days after denervation, we isolated single muscle fibers from control and denervated soleus and EDL muscles via collagenase digestion (Figure 2A, C). Separated fibers were manually isolated, digested using the protease trypsin, and analyzed using a 4 h LC-MS/MS gradient (Figure 2E). This approach is complicated by the fact that the low amount of proteins in individual

muscle fibers compromises in-depth proteome analysis.



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To increase the number of protein hits, we pooled ~150 muscle fibers from several muscles and analyzed the samples using a 4 h gradient as well, to ensure similar retention times for the peptides in the single fiber analysis and peptides in the reference dataset (7, 38) (Figure 2B, D, F). Here each peptide is represented by its retention time, accurate mass and associated MS/MS fragmentation spectra (Figure S-1A). In total, the muscle reference dataset contains 47,032 identified peptides, along with the MS/MS spectra and retention times. The comparisons with the single fiber analysis were performed via the “match between runs” option of MaxQuant software (Table S-2). The overlap to an earlier study describing the proteome of different single muscle fiber types (7) revealed an overlap of ~900 proteins (Figure S-1B), indicating similar matches between both datasets. It is important to note that MyHC isoforms have a very high sequence similarity, thus correct peptide assignment to specific MyHC isoforms based on shotgun proteomics is challenging. To avoid any misinterpretation, peptide assignment was restricted to unique MyHC peptides, and the relative abundance of each MyHC isoform in individual fibers was calculated relative to the sum of the intensity of the four MyHC isoforms (MYH7, MYH2, MYH4, MYH1) in each fiber.


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Figure 2: Experimental workflow for the analysis of single muscle fibers

(A) Mouse hind leg muscles were denervated by section of the sciatic nerve and the soleus and EDL of the control and denervated legs were isolated 7 days after denervation. (B) The muscle peptide reference dataset was generated by collagenase digestion of seven different muscles. (C) Soleus and EDL muscles were digested with collagenase and ~50 single fibers were isolated from control and denervated soleus fibers, respectively. Similarly, ~25 single fibers were isolated from the EDL. (D) Approximately 150 fibers were collected for each muscle to establish a peptide reference dataset representing seven muscles. (E) Samples were assessed using a 4 h LC-MS/MS gradient and analyzed with the “match between runs” option implemented in MaxQuant software. (F) Fiber types were assigned to single muscle fibers based on MyHC composition and pooled accordingly. Fibers from control and denervated muscles were grouped and directly compared.

Similar to earlier studies (7), fibers that expressed a high level of a specific MyHC isoform were assembled to that specific MyHC group, allowing direct comparison of different fiber types under control and denervated conditions (Figure 2G). For quantitative analysis of the whole dataset, we used unique and razor peptides and identified 23,432 peptides corresponding to 2,270 proteins across all fibers from the soleus and EDL, with a FDR below 1% (Figure 3A, Figure S-2A and Table S-2). Raw data analysis without utilizing the reference dataset resulted in ~25% fewer protein identifications, indicating improved protein identification (Figure 3A). Consistent with previous reports, ~48% of the investigated soleus fibers were MYH7positive and classified as type I fibers, ~44% were MYH2-positive (type IIa), and ~8% had mixed MYH7/MYH2 or MYH2/MYH1 compositions (24, 25) (Figure 3B). As a consequence of denervation, we observed a slight reduction in


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Figure 3: Quantitative analysis of MyHC expression during muscular atrophy

(A) Number of proteins identified and quantified from soleus and EDL fibers. Integration of the peptide reference dataset enhanced protein identification by ~25%. (B) Fiber type profiling of control and (C) denervated soleus muscles. (D) Differential expression of MyHC isoforms in different fiber types, * significantly regulated (FDR < 5%). (E) Principal component analysis (PCA) separated type I from type IIa fibers, and denervated from control fibers. (F) Average number of proteins quantified per single fiber in type I, IIa and IIb fibers under control and denervated conditions. (G, H) Cumulative intensity distribution of proteins in type I (G) and (H) type IIa fibers. Ident.: Identified, Quant.: Quantified

type IIa fibers and increase in hybrid MYH7/MYH2 or MYH2/MYH1 fibers in the soleus (Figure 3C). Analysis of EDL fibers substantiated the high number of type IIb fibers under normal conditions (Figure S-2B) (23), and we observed a slight increase in the proportion of fibers expressing MYH1 (type IIx) in the EDL after denervation (Figure S-2C). Reduced neuronal innervation converts slow fibers into faster fibers by altering the MyHC composition from MYH7 to faster isoforms, including MYH2, MYH1 and MYH4 (2, 39, 40). Conversely, fast fibers tend to shift to express the slower MyHC isoforms (41). After denervation, type I and IIa fibers showed a significant upregulation of the fast MYH4 isoform, suggesting initiation of a fiber type transition in the soleus (Figure 3D). Conversely, denervated type IIb fibers showed enhanced expression of MYH7. In addition, we also observed increased levels of the embryonic myosin MYH3 in denervated type IIa and IIb fibers (Figure 3D) (42, 43). However, since MyHC have relatively long half-lives of ~ 28 days in rabbits and ~54 days in rats (44, 45), a slight increase in MYH4 expression in the soleus might not be adequate to induce a marked fiber type transition at this early time point after denervation (46, 47). Collectively, our quantitative proteomic analyses of single fibers


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reflected the known fiber type compositions of the soleus and EDL muscle and enabled comparison of fiber types with the same MyHC expression profile and metabolic activity during muscular atrophy.

Systematic fiber type clustering reveals early remodeling activities during atrophy

To determine whether the loss of neuronal stimulation has distinct effects on the proteomes of individual fibers, we performed principle component analysis (PCA) and observed clear separation of control and denervated fibers, as well as type I and type IIa fibers, by the first two components (Figure 3E). Similarly, PCA also showed clear separation of the control and denervated type IIb fibers in the EDL (Figure S-2D). On average, we quantified ~7,000 peptides, resulting in identification of ~600 proteins per single muscle fiber (Figure S-2E, Table S-2). We observed a clear reduction in the number of protein hits in all denervated soleus fibers compared to control fibers (Figure 3F). Next, we ranked all proteins according to their IBAQ intensity and calculated their cumulative abundance (Figure 3G-H; Figure S-2F). In skeletal muscle cells a relatively small number of myofibrillar proteins account for the majority of the protein content (48, 49). For example, the four most abundant proteins - two MyHCs, titin and actin - account for ~50% of the entire protein content in each muscle fiber. Hence, the low protein concentration, high dynamic range, and analysis of isolated muscle fibers without other cell types and extracellular matrix contributed to the low protein identification rate for single muscle fibers (Figure 3G-H). Similar to the quantitative analysis of intact muscles, we performed another set of two sided t-tests with FDR multiple testing correction (Table S-2). After sorting fibers with the same MyHC expression into distinct groups (Figure 2G), we quantified 1,762 proteins and observed significant differential regulation of ~35% of the proteins in each fiber type.



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Loss of neuronal innervation induced downregulation of the highest number of proteins in type I and type IIa fibers in the soleus, whereas equal numbers of proteins were up- and downregulated in the - IIb fibers of the EDL after denervation (Figure 4A-C). Comparison of all differentially regulated proteins revealed four proteins, the ribosomal protein RPL18, the myosin light chain 12a (MYL12A), the major vault protein (MVP), and the pre-B-cell leukemia transcription factor-interacting protein (PBXIP1) - were commonly upregulated in all fiber types (Table S-3). PBXIP1 interacts with the homeodomain protein PBX1 and inhibits DNA binding of the PBX1-HOX complex, a known regulator of gene expression in hematopoietic cell populations (50). Since we observed a general increase in protein expression in all muscle fibers, PBXIP1 may also function as a repressor of PBX1 during muscle atrophy. The exclusive upregulation of 45 proteins in type IIb fibers from the EDL substantiates the different response of these muscle fibers to loss of neuronal innervation compared to soleus fibers. In addition, 43 proteins were downregulated in all tested fibers (Table S-3). For example, the monophosphate deaminase AMPD1, the E3 ubiquitin ligase TRIM72 (also known as MG53), and the histone-lysine N-methyltransferase SMYD1 were


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Figure 4: Fiber type-dependent protein changes after denervation



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(A) Differentially regulated proteins in type I and type IIa fibers of the soleus were identified by plotting the -log10 P-values and log2 fold changes in label-free protein intensities. Volcano plot with a 5% FDR curve identified 193 differentially regulated proteins in type I soleus fibers, (B) 230 in type IIa soleus fibers and (C) 264 in type IIb EDL fibers. (D) Scatter plot of log2 den/ctrl ratios for type I and IIa fibers. The inset illustrates the opposing regulation of ATP2A2 in type I and IIa fibers. Enlarged data points indicate proteins that were significantly regulated in at least one fiber type. Red and blue areas indicate the 1.5 foldchange cut-off. (E) Selected marker proteins with specific expression in slow (yellow) or fast (green) fibers and their protein rank within the whole protein distribution (upper panels) and box plot of log2 den/ctrl ratios of the same slow and fast marker proteins in type I (left) and type IIa fibers (lower panel). (F) Immunohistochemical stainings for the indicated proteins on muscle cryo-sections from control and denervated soleus muscle. Fiber types were determined by staining of consecutive sections with MYH7 antibodies. “X” marks identical fibers in serial cross-sections. Box plots show relative fluorescence intensity of individual fibers (n=10). P-values were calculated using Whitney-Mann U test. Scale bars = 50 µm. (G) Immunoblot analysis of CRYAB levels in type I single muscle fibers. Actin levels detected in the Stain-Free acquisition of the blot serve as loading control.

downregulated equally upon denervation in type I, IIa and IIb fibers (Table S-3). AMPD1 plays an important role in energy production by converting AMP to IMP within the purine nucleotide cycle; downregulation of AMPD1 may reflect reduced energy production during muscular atrophy. TRIM72 is mainly expressed in type I fibers and plays a central role in membrane repair, acts as a sensor for protein oxidation and is a negative regulator of myogenesis (51, 52). While other TRIMs, including TRIM25 and TRIM63 (MURF1), were


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upregulated during muscular atrophy, downregulation of TRIM72 after atrophy suggests a yet unknown function for this E3 ubiquitin ligase in type I fibers.

Type I and type IIa fibers of the soleus exhibit opposing protein changes during muscle atrophy In addition to detection of proteins that were similarly regulated after denervation, we also detected proteins that were adversatively regulated in different fiber types. In order to visualize this effect, we plotted the log2 control/denervated ratios for type I and type IIa fibers (Figure 4D). We detected 34 proteins that were exclusively and significantly regulated in type I fibers and 84 proteins with regulation in type IIa fibers (cutoff < 1.5-fold, P-value < 5% FDR; Figure 4D). For example, the slow sarcoplasmic/endoplasmic reticulum calcium ATP2A2 (SERCA2) was significantly downregulated in type I fibers and upregulated in type IIa soleus fibers. Importantly, we detected no change in ATP2A2 protein expression in analysis of the intact soleus tissue (Figure 4D insets; Table S-1 and S-2), suggesting that the opposing responses of ATP2A2 in different fiber types during muscular atrophy made it impossible to detect this differential regulation of ATP2A2 in the intact soleus. In addition, a selection of fiber type-specific protein regulations are listed, with the corresponding fold-changes and Pvalues, in Table 1. Next, we focused our analysis on eight known slow and fast marker proteins (2) and visualized their abundance within the proteome of type I and type IIa fibers (Figure 4E, upper panels). The boxplot analysis showed enhanced proteins levels of fast marker proteins (green box) in type I fibers after denervation compared to the whole dataset (P < 5E-2) and slow marker proteins (yellow box, P < 5E-2; Figure 4E, left panel). Conversely, this tendency was not detectable for the same slow and fast marker proteins compared to the entire ratio distribution of type IIa fibers (right side). To substantiate our



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mass spectrometric data we selected ATP2A2 and CRYAB for further validation and performed immunostainings and on control and denervated soleus muscles and single fiber western blots (Figure 4F-G). CRYAB is an important chaperone for muscle maintenance and correct folding of sarcomeric proteins such as actin, desmin and titin (53, 54). Its downregulation could make fibers more sensitive to denervation-induced atrophy.

Table 1: Selection of fiber specific regulated proteins, which are not detectable in whole muscles. Statistical significance (+) is based on permutation based multiple testing

correction with 5% FDR. Den/Ctrl Type I Protein

ATP2A2

Protein name

SR calcium ATPase 2

ARL6IP5 PRA1 family protein 3

Den/Ctrl Type IIa

Type IIa Ctrl/Type I Ctrl

-Log10

Log2 Signifi-

-Log10

Log2 Signifi-

-Log10

Log2 Signifi-

P-value

Ratio

cant

P-value

Ratio

cant

P-value

Ratio

cant

5.875

-1.23

+

1.249

1.17

+

2.662

-0.91

+

0.176

-0.11

3.654

1.09

+

14.140

-5.65

+

CANX

Calnexin

0.008

0.003

3.347

1.01

+

4.838

-0.97

+

TNNC1

Troponin C, slow

0.033

-0.02

2.354

1.49

+

14.954

-5.11

+

TNNT1

Troponin T, slow

0.775

0.23

6.955

1.86

+

27.010

-5.88

+

MYL6B

Myosin light chain 6B

0.009

-0.005

2.580

1.86

+

9.593

-3.95

+

0.476

-0.44

1.379

0.96

+

0.902

-0.53

ABLIM1

Actin-binding LIM protein 1

Muscle fibers have several metabolic pathways to supply the contractile machinery with ATP in order to produce sufficient kinetic energy for motion. For instance, creatine kinase and glycolytic enzymes are the key players in generation of energy in the form of ATP for rapid muscle contraction. Long term energy supply is mediated by the oxidation of acetylCoA via the tricarboxylic acid cycle (TCA) and oxidative phosphorylation (OXPHOS) complexes. Once sugar and glycogen stores are depleted, muscles can also metabolize proteins and fatty acids to produce energy. Each fiber type has distinct physiological adaption 24 ACS Paragon Plus Environment

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and is therefore associated with a specific composition of metabolic enzymes. Our quantitative mass spectrometric analysis confirmed the fiber type-specific distribution of enzymes associated with glycolysis, fatty acid metabolism and oxidative phosphorylation (Table S-2), in agreement with previous reports (55, 56). Overlapping our datasets with the Mitocarta database revealed more than 200 mitochondrial proteins were detected across all fiber types, and indicated a slight decrease in mitochondrial proteins in type IIa fibers of soleus and type IIb fibers of the EDL after denervation compared to type I fibers from the soleus after denervation (Figure 5A). A volcano plot analysis of type I and type IIa fibers from control and denervated soleus also revealed higher levels of mitochondrial proteins in type IIa fibers under normal conditions, and significant downregulation of most of these proteins after denervation (P < 0.0; Figure 5B, C). This could suggest soleus type IIa fibers have a higher sensitivity to atrophy. However, proteins such as isocitrate dehydrogenases (IDH3B and IDH3G), D-betahydroxybutyrate dehydrogenase (BDH1) and the mitochondrial ribosomal protein S36 (MRPS36) were still more abundant in type IIa fibers than in type I fibers after denervation (Figure 5C). The TCA cycle protein NAD-dependent isocitrate dehydrogenase 3 (IDH3) is involved in oxidative decarboxylation of isocitrate into α-ketoglutarate and consists of three subunits (IDH3A, IDH3B, IDH3G). Similarly to a previous study on single muscle fibers, we observed higher levels of IDH3 in type IIa control muscle fibers and higher levels of IDH2 in type I- control muscle fibers (Figure 5B) (7). Although all three IDH3 subunits were significantly downregulated after denervation, the abundance of the IDH3A subunit decreased more than that of IDH3B/G (Figure 5B; Figure S-3C, Table S-2).



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Figure 5: Systematic analysis of protein changes reveal functional characterization of denervated muscle fibers

(A) Box plot of the log2 den/ctrl ratio distribution of mitochondrial proteins. (B) Volcano plots comparing protein fold changes in mitochondrial proteins between type I and IIa fibers in control soleus fibers. Green and yellow circles represent the 44 significantly differentially expressed mitochondrial proteins between type I and type IIa fibers (permutation-based FDR cutoff of 5%). (C) Volcano plot for 215 mitochondrial proteins in type I and type IIa fibers after denervation. (D) Box plot analysis of OXPHOS complexes for type I, IIa and IIb fibers between control and denervated fibers. (E) Changes in glycolysis pathway proteins in type I, IIa and IIb fibers. (F) Box plots of log2 den/ctrl ratios of proteasomal subunits and (G) graphical representation of proteasomal subunits and their changes after denervation.

In addition to its function as an important TCA cycle enzyme, IDH3A has also been associated with cancer; high levels of IDH3A may promote tumor growth by increasing the levels of α-ketoglutarate and HIF1 (57). Since mitochondrial proteins are not uniformly regulated after denervation, we hypothesize that differential regulation of subsets of mitochondrial proteins may modulate the metabolic activity of specific muscle fibers during atrophy. Hence, single fiber proteomics will contribute to decipher the heterogeneous changes in mitochondrial proteins in fibers expressing different MyHC isoforms. The mitochondrial contact site and cristae organizing system (MICOS) is another complex that exhibited heterogeneous regulation. Although most members were significantly downregulated after denervation, metaxin-2 (MTX2) was upregulated in type I and type IIa fibers (Figure S-3D, Table S-2). The MICOS complex is required for formation of cristae and modulates the activity of respiratory complexes. MTX2 may potentially maintain protein



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import into mitochondria, since it is localized at the outer mitochondrial membrane and associates with the mitochondrial intermembrane space bridging complex (MIB). A boxplot analysis of the OXPHOS complexes (I-V) revealed downregulation of complex III and IV in type IIa and IIb fibers after denervation, whereas these proteins were less affected in type I fibers (Figure 5D; Table S-4). Conversely, the hexokinase 2 (HK2), the first enzyme of glycolysis, was significantly downregulated in type I and IIa fibers - but not type IIb fibers - after denervation, indicating the importance of glycolysis in fast type IIb fibers (Figure 5E). However, the rate limiting enzyme of glycolysis, phosphofructokinase (PFKM), was significantly downregulated after denervation in all three fiber types (Figure 5E). The proteasome was another heterogeneously regulated cellular compartment; we quantified 29 proteasomal subunits. The boxplot analysis substantiated downregulation of several proteasomal subunits in type I fibers after denervation compared to type IIa and IIb fibers (P < 10E-2; Figure 5F-G; Table S-4). For example, the subunits PSMA6 and PSMD2 were downregulated in denervated type I fibers, whereas eight proteasomal subunits were significantly upregulated in denervated type IIb fibers compared to control fibers.

Loss of neuronal innervation induces fiber type-specific dysregulation of calcium homeostasis Next, we assessed the fiber type-specific response of proteins involved in calcium homeostasis. First, we ranked all calcium-associated proteins based on their fiber typespecific label free protein intensity (LFQ) to assess their relative abundance within type I and IIa fibers (Figure 6A-B). For example, the tropomyosin isoforms TPM1 and TPM2 were upregulated in type IIa fibers, but not in type I fibers after denervation. Troponin C is a calcium binding protein that interacts with troponin I (TNNI) and troponin T (TNNT) to


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regulate skeletal muscle cell contraction (58). Each troponin subunit isoform is expressed in a fiber type-specific manner, and in agreement with earlier reports, we confirmed troponin C (TNNC1) was expressed in type I fibers in slow skeletal and cardiac muscle. Accordingly, troponin C2 (TNNC2), the skeletal muscle-specific isoform, was present at higher levels in type IIa fibers compared to type I fibers (Figure S-3A-B). After denervation, we noticed significant upregulation of ‘slow’ TNNC1 in type IIa fibers and enhanced levels of TNNC2 in type I and type IIa fibers. Similarly, TNNT1 was also upregulated in type I fibers and the corresponding ‘fast’ TNNT3 isoform was upregulated in both type I and IIa fibers after denervation. In contrast, TNNI1, which inhibits troponin complex formation, was significantly downregulated in type I fibers after denervation (Figure 6A-B; Figure S-3F). Although the physiological relevance of these remodeling processes needs to be elucidated in future experiments, our experimental approach shows the potential to study fiber type-specific protein changes.

Figure 6: Calcium-associated proteins are upregulated in type IIa fibers after denervation (A+B) Ranked LFQ protein intensities of calcium signaling-related proteins in control fibers indicating their abundance and differential expression after denervation (A) in type I and (B) in type IIa fibers. 29 ACS Paragon Plus Environment


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DISCUSSION The analysis of muscles as a whole always reflects a mixture of different fibers, extracellular proteins, cell types and blood vessels. Moreover, an additional layer of complexity arises from the varied responses of each fiber type to environmental influences; these responses are modulated by neuronal innervation, metabolic activity and the MyHC expression profile (19, 41). In order to follow protein dynamics in individual fibers during muscular atrophy, we employed a mouse model involving section of the sciatic nerve and performed single muscle fiber proteomic analysis of individual slow type I, intermediate type IIa and fast IIb fibers from control and denervated muscles seven days after denervation. Since we focused on a relatively early time point after denervation (7 days), we did not detect marked fiber type switching based on complete MyHC transitions. However, we quantified an initial increase in the proportions of MYH1- and mixed MYH4/MYH1-positive fibers (~12%) in the EDL, indicating an early transition from IIb- to IIx-positive fibers. Likewise, analysis of type I fibers from the soleus revealed expression of MYH4 was significantly enhanced, from ~1% in control fibers to ~4% in 80% of all denervated type I fibers (Mann-Whitney test: P = 1E-06). This upregulation of MHY4 may be restricted to a specific subset of fibers, since MYH4 was undetectable in ~20% of denervated soleus type I fibers. Earlier studies showed slow type I fibers can be subdivided into primary and secondary fibers, which express different sets of MyHC isoforms during embryonic and postnatal development. For instance, secondary fibers are MYH2-positive during embryonic development and later switch to type I (59). Conversely, primary fibers exclusively express MYH7 and are potentially more resistant to a transition to faster fibers. According to


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previous studies it might be possible that the ~20% of type I fibers in which MYH4 was undetectable at 7 days after denervation represent primary fibers. However, since MyHC proteins have a rather long half-life of ~30 to 50 days (44, 45), it would be interesting to determine the kinetic profiles of MyHC expression in individual fibers from a diverse set of skeletal muscles. This would help to unravel the distribution of primary and secondary fibers and possibly reveal how, depending on their localization, the MyHC profiles of these fibers are altered during catabolic conditions. Although we were able to quantify fewer proteins compared to analysis of whole muscle lysates, our single fiber proteomics approach allowed us to detect 60 differentially regulated proteins which were not observable without separating the individual fiber types. Earlier studies found the rate of protein synthesis in the intact EDL muscle increases at 7-10 days after denervation, with only minor muscular atrophy evident at this time-point (60). This anabolic effect in the EDL can be explained by its greater extent of passive stretch compared to the slow soleus muscle, which induces protein synthesis even in the absence of neuronal innervation (61). Accordingly, we demonstrated that the type IIb fibers of the EDL contained the highest number of upregulated proteins after denervation. We also observed enhanced expression of eight proteasomal subunits in type IIb fibers, and it seems reasonable to assume that increased levels of proteasomal subunits are required to balance enhanced protein synthesis in response to denervation. An important pathway to regulate protein turnover is the activation of the AKT-mTOR pathway (62, 63). Hence, it might be interesting to investigate whether elevated protein degradation is also associated with a fiber typespecific activation of the mTOR pathway and its downstream targets. Although not feasible yet, phosphoproteomics on muscle fibers might contribute to dissect the activity of signaling pathways in fibers with the same MyHC expression and metabolic activity in the future.



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Uptake of proteins into lysosomes is another pathway of protein degradation. These digestive organelles contain several proteases, such as cysteine and aspartyl proteases. We found enhanced levels of cathepsin B, D and L1 proteases in type I and type IIb fibers, whereas expression of these proteases did not change in type IIa fibers (Table S-2). Thus, it is possible that specific adaptions to the lysosomal protein degradation machinery occur after denervation in muscle fibers with different metabolic and contractile activities. Mitochondria are crucial for energy production and several metabolic pathways are integrated within this organelle (Figure S-3E). Consistent with previous reports, we found type IIa fibers had the highest abundance of mitochondria, and our quantitative analysis showed most mitochondrial proteins were downregulated after denervation. Accordingly, we also observed reduced levels of several key enzymes in the TCA cycle, β-oxidation and OXPHOS complexes in all fiber types. Moreover, our study revealed that most enzymes involved in glycogen hydrolysis were more abundant in type IIb fibers than in IIa and I fibers, confirming the finding that fast fibers preferentially use glycogen as a source to provide glucose-6-phosphate as an initial metabolite for carbohydrate metabolism (Table S2) (64). Unsurprisingly, most glycogenolytic enzymes were also downregulated in all fiber types after denervation and the largest numbers in protein reduction were observed in type IIb fibers. This suggests that the supply of energy from glycogen metabolism is disrupted to a higher extent in fast fibers than in other fibers after denervation (Figure S-3G, Table S-2). The hexokinase HK2 phosphorylates glucose and represents a functional association between glycolysis and the OXPHOS complexes (65). Moreover, HK2 inhibits apoptosis by reducing the permeability of the mitochondria to Bax and Bak, and the interaction of HK2 with the fructose-2, 6-bisphosphatase TIGAR enhances HK2 activity (66, 67). Although the TIGARHK2 interplay has been described under hypoxic conditions and we could not quantify


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TIGAR in soleus fibers, it is possible that HK2 connects energy availability and mitochondrial activity during catabolic conditions.

Changes in calcium homeostasis in atrophying muscle Skeletal muscles have several transporters and Ca2+ binding proteins to rapidly modulate the Ca2+ ion concentrations in their cytoplasm and cellular compartments including the sarcoplasmic reticulum (SR), mitochondria and T-tubules. The most abundant calcium transporters are sarcoplasmic/endoplasmic reticulum calcium ATPases and our quantitative analysis revealed opposing regulation of ATP2A2 in type I and IIa fibers, which initially prevented detection of fiber type-specific regulation of this protein in the intact soleus. Consistently, a previous study of denervated mouse soleus muscle also showed no changes in ATP2A2 protein expression within the first two weeks of denervation based on western blot analysis (68). It is possible that type IIa fibers modify calcium homeostasis by upregulating ATP2A2 in response to the increased Ca2+ levels after denervation. This remodeling was also reflected by enhanced levels of several calcium binding proteins with EF-hand domains in type IIa fibers, including troponin C (slow and fast), calmodulin, parvalbumin and calumenin (Table S-2). In particular, upregulation of the sarcoplasmic reticulum Ca2+ binding protein calsequestrin-1 (CASQ1) may help to reduce the cytoplasmic Ca2+ concentration by keeping the ions in the sarcoplasmic reticulum. We also observed significantly enhanced levels of calumenin (CALU), another recently identified calcium binding protein, in type IIa fibers. Calumenin is involved in Ca2+ uptake into the SR, and overexpression of calumenin reduces Ca2+ uptake by inhibiting ATP2A2 (69). Hence, enhanced calumenin expression may reflect a compensatory response to increased ATP2A2 levels in type IIa fibers. Another regulator of ER protein secretion is calnexin (CNX), a calcium-dependent membrane protein that is also present in the SR membrane (70-72). CNX functions together with calreticulin (CALR) to



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regulate glycoprotein folding and assembly (73). Moreover, calnexin also participates in the ER protein quality control system by preventing non-native proteins from being secreted from the ER (72, 74, 75). We found calnexin was upregulated in type IIa and IIb fibers - but not in type I fibers - after denervation (Table 1, Table S-1). The expression of soluble calreticulin, which exerts similar functions as calnexin (75), was not altered after denervation. Hence, we suggest that calnexin may more frequently assist glycoprotein folding and ER quality control during catabolic conditions than calreticulin. Overall, this single muscle fiber proteomics approach reveals a number of quantitative differences between different fiber types and demonstrates the heterogeneous response of muscle fibers with different MyHC expression profiles and metabolic activity. Encouragingly, this single fiber proteomics approach could be used to investigate the response of different muscle fiber types to any perturbation, including metabolic syndromes and neuronal diseases.

ACKNOWLEDGEMENTS We acknowledge Sylvia Jeratsch for technical assistance and Kerstin Selbach for helpful comments on this manuscript. This work was supported by the Cologne Cluster of Excellence in Cellular Stress Responses in Aging-associated Diseases (CECAD).

CONFLICT OF INTEREST DISCLOSURE The authors declare no competing financial interest. SUPPLEMENTAL INFORMATION Figure S-1: Peptide matching between single fiber samples and the peptide reference dataset Figure S-2: Summary of protein changes in different fiber types after denervation Figure S-3: Functional characterization of fiber types after denervation


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Table S-1: Proteomic dataset of whole muscles (EDL, GAST, Soleus, TA) Sheet 1 Differentially expressed proteins in four different muscles following denervation Sheet2 Protein quantification data for EDL, GAST, Soleus, TA, including LFQ intensities, number of peptides and sequence coverage Sheet 3 Sequences of identified peptides in control and denervated muscles

Table S-2: Proteomic Profiling of single fibers from soleus and EDL muscle upon short-term denervation Sheet 1 Fiber classification based on relative abundance of MyHC isoforms. Sheet 2 Differentially expressed proteins in different fiber type groups. Sheet 3 Protein quantification of single fibers and the muscle peptide reference dataset, including LFQ intensity, number of peptides and sequence coverage. Sheet 4 Sequences of identified peptides in single muscle fibers and the muscle peptide reference dataset. Table S-3: Overlap of regulated proteins between type I, IIa and IIb fibers Sheet 1: Commonly up regulated proteins Sheet 2: Commonly down regulated proteins Sheet 3: Proteins exclusively regulated in type I fibers Sheet 4: Proteins exclusively regulated in type IIa fibers Sheet 5: Proteins exclusively regulated in type IIb fibers Table S-4: Proteomic changes of selected compartments Sheet 1 Protein expression levels of Complex I-V proteins. Sheet 2 Protein expression levels of proteasomal subunits quantified based on unique peptides.



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Sheet 3 Protein expression levels of calcium-related proteins.

For Table of Contents only

Uncategorized References 1. Allen, D. G.; Lamb, G. D.; Westerblad, H., Skeletal muscle fatigue: cellular mechanisms. Physiol Rev 2008, 88, (1), 287-332. 2. Schiaffino, S.; Reggiani, C., Fiber types in mammalian skeletal muscles. Physiol Rev 2011, 91, (4), 1447-531. 3. Fry, A. C.; Allemeier, C. A.; Staron, R. S., Correlation between percentage fiber type area and myosin heavy chain content in human skeletal muscle. Eur J Appl Physiol Occup Physiol 1994, 68, (3), 246-51. 4. Murgia, M.; Toniolo, L.; Nagaraj, N.; Ciciliot, S.; Vindigni, V.; Schiaffino, S.; Reggiani, C.; Mann, M., Single Muscle Fiber Proteomics Reveals Fiber-Type-Specific Features of Human Muscle Aging. Cell Rep 2017, 19, (11), 2396-2409. 5. Lecker, S. H.; Jagoe, R. T.; Gilbert, A.; Gomes, M.; Baracos, V.; Bailey, J.; Price, S. R.; Mitch, W. E.; Goldberg, A. L., Multiple types of skeletal muscle atrophy involve a common program of changes in gene expression. FASEB J 2004, 18, (1), 39-51. 6. Spangenburg, E. E.; Booth, F. W., Molecular regulation of individual skeletal muscle fibre types. Acta Physiol Scand 2003, 178, (4), 413-24. 7. Murgia, M.; Nagaraj, N.; Deshmukh, A. S.; Zeiler, M.; Cancellara, P.; Moretti, I.; Reggiani, C.; Schiaffino, S.; Mann, M., Single muscle fiber proteomics reveals unexpected mitochondrial specialization. EMBO reports 2015, 16, (3), 387-395.


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