by P21 (**Figure 2**; **Table 1**). Thin filament lengths determined by F-actin breadth from the Z-line to the H-zone paralleled those determined by Tmod1 localization in both neonatal GAS and TA muscles at all time-points (**Table 1**). Nebulin M1M2M3 distances from the Z-line remained constant at ∼0.94μm in both neonatal GAS and TA (**Figure 2**; **Table 1**), consistent with the nebulin-coated proximal segment of the thin filament having a fixed length, regardless of the length of the nebulin-free distal segment (Gokhin and Fowler, 2013).

#### **Table 1 | Thin filament lengths determined by Distributed Deconvolution analysis of fluorescence images.**

*Length values (all in µm) correspond to the breadth of phalloidin staining and the distances of Tmod and nebulin M1M2M3 from the Z-line. Data are shown mean* ± *SD for n* ≥ *100 myofibrils/time-point for Tmod and phalloidin, n* ≥ *50 myofibrils/time-point for nebulin M1M2M3. \*p < 0.01 with respect to the previous time-point.*

Next, to examine changes in thin filament length during skeletal muscle aging, we further used confocal fluorescence microscopy and Distributed Deconvolution analysis to compare thin filament lengths in 2-mo-old vs. 2-yr-old mice. Aged muscles showed no evidence of gross myofibril disorganization or deterioration, as evidenced by striated Tmod1, nebulin M1M2M3, F-actin, and α-actinin staining (data not shown). In the TA and EDL, no statistically significant differences in thin filament lengths were observed between 2-mo-old vs. 2-yr-old muscles (∼1.11μm in both muscles at both time points) (**Figure 3**; **Table 1**). However, in the soleus, thin filament length decreased from ∼1.20μm in 2-mo-old muscles to ∼1.11μm in 2-yr-old muscles (**Figure 3**; **Table 1**). As observed with our measurements of thin filament lengths in neonatal muscles, thin filament lengths determined by F-actin breadth from the Z-line to the H-zone paralleled those determined by Tmod1 localization (**Table 1**), and nebulin M1M2M3 distances from the Z-line remained constant at ∼0.95μm (**Figure 3**; **Table 1**). Moreover, thin filament lengths in the TA muscles of 2-mo-old BALB/cBy mice were identical to thin filament lengths in the TA muscles of P21 C57Bl/6J mice (**Table 1**; also compare **Figure 2** and **Figure 3**), indicating that thin filament lengths do not change between P21 and 2 months of age, and that lengths are not mouse strain-dependent. Thin filament lengths in adult mice in these strains also agree with lengths previously reported in adult mixed-background FVB/N:129/SvJ:C57Bl/6 mice (Gokhin et al., 2010).

### **DISCUSSION**

This study provides the first evidence that skeletal muscle thin filaments undergo adaptive remodeling to alter their lengths during the mouse lifespan, implying age-dependent alterations in sarcomere length-tension relationships and optimum sarcomere length ranges. What molecular mechanisms might drive these changes in thin filament lengths? When examining neonatal muscles, we observed thin filament shortening between P7 and P21 in both the TA and GAS (**Figure 2**). These changes are mostly associated with developmental transitions away from expression of embryonic and neonatal MHC isoforms and toward expression of adult type-II MHC isoforms (Allen and Leinwand, 2001;

from the Z-line in TA, EDL, and soleus muscles from 2-mo-old to 2-yr-old mice. Tmod distance reflects the variable position of the thin filament pointed end with respect to the Z-line, indicative of variations in length, while nebulin M1M2M3 distance reflects the relatively constant position of the proximal/distal segment boundary of the thin filament with respect to the Z-line (Gokhin and Fowler, 2013). Tmod distances reflect pooled Tmod1 and Tmod4 distances. Error bars reflect mean ± *SD* for *n* ≥ 100 myofibrils/time-point for Tmod, *n* ≥ 50 myofibrils/time-point for nebulin M1M2M3. ∗*p <* 0*.*01 with respect to the previous time-point.

Agbulut et al., 2003; Gokhin et al., 2008). The presence of type-II MHC isoforms is, in turn, associated with shorter thin filament lengths (Castillo et al., 2009; Gokhin et al., 2010, 2012). This suggests a model in which immature mouse muscles, expressing predominantly embryonic and neonatal MHCs, are all initially specified with "generic" thin filament lengths during the terminal stages of myofibril assembly, despite these muscles' functional differences in adult mice (Allen and Leinwand, 2001; Agbulut et al., 2003; Gokhin et al., 2008). Then, as the various muscles adopt their characteristic use profiles and MHC isoform distributions during postnatal acquisition of adult movement patterns, thin filament lengths may gradually diversify, ultimately leading to the multitude of muscle-specific thin filament lengths observed in adult mice (i.e., longer thin filaments characteristic of slower fiber types and shorter filaments characteristic of faster fiber types) (Gokhin et al., 2010, 2014). Simultaneously, postnatal changes in tropomyosin and troponin isoform expression (Amphlett et al., 1976; Briggs et al., 1990) may also be involved in regulating thin filament lengths during postnatal skeletal muscle development. Evidence supporting this possibility is the fact that a nemaline myopathy-causing tropomyosin mutation can cause markedly shorter thin filaments *in vivo* (Ochala et al., 2012), and that both tropomyosin and troponin have been shown to enhance actin filament pointed-end stability *in vitro* (Broschat et al., 1989; Broschat, 1990; Weigt et al., 1990).

When comparing 2-mo-old and 2-yr-old muscles, we observed no statistically significant changes in thin filament length in the TA or EDL, but a marked decrease in thin filament length was observed in the 2-yr-old soleus (**Figure 3**). The fact that thin filament shortening during aging is restricted to the soleus muscle implies a mechanism specific to heavily recruited, slow-twitch muscles. Such a mechanism most likely does not involve shifts toward expression of type-II MHC isoforms during aging, unlike the MHC isoform shifts described above for postnatal muscle development, because studies of rat soleus have shown continued enrichment of type-I MHC with decreases in type-II MHC during aging (Butler-Browne and Whalen, 1984; Larsson et al., 1995). Indeed, an increase in type-I MHC with concomitant thin filament shortening is not consistent with a causative effect of MHC isoforms in regulating thin filament lengths. Other molecules whose alterations might directly induce slow-muscle-specific thin filament shortening in aging are tropomyosin (an actin sidebinding and stabilizing protein), whose α isoforms are decreased in aged muscle (Gelfi et al., 2006; O'Connell et al., 2007), as well as ADF/cofilin (an actin-severing protein) and profilin (an actin monomer-sequestering protein), whose levels are increased in degenerating muscle (Nagaoka et al., 1996). Moreover, slowmuscle-specific changes in thin filament protein phosphorylation, glycosylation, or other posttranslational modifications might also contribute to slow-muscle-specific thin filament shortening in aging. Understanding the contributions and interplay of these molecules and pathways to thin filament shortening in aging requires further studies.

An alternative mechanism for slow-muscle-specific thin filament shortening in aging involves preferential induction of proteolytic pathways in slow-twitch muscles (Sultan et al., 2001), leading to breakdown of thin filament-associated proteins and reduced F-actin stability. Such proteolytic pathways might be specifically induced in old muscle, or might be ongoing in heavily recruited muscles and begin in mid-life. A number of proteolytic pathways have been implicated in skeletal muscle aging and aging-related sarcopenia, including activation of intracellular proteases (calpains and caspases), the ubiquitin-proteasome system, the autophagy-lysosome system, and reactive oxygen species (for reviews, see Fulle et al., 2004; Dargelos et al., 2008; Ryall et al., 2008; Combaret et al., 2009). Activation of m-calpain can lead to Tmod proteolysis that is more widespread in slow muscle in mouse models of Duchenne muscular dystrophy, but this mechanism results in actin subunit addition onto pointed ends and resultant thin filament elongation, and not thin filament shortening (Gokhin et al., 2014). More likely targets whose proteolysis could lead to shorter thin filaments are actin filament side-binding proteins such as nebulin and tropomyosin, which stabilize thin filaments to prevent aberrant thin filament shortening (Bang et al., 2006; Ottenheijm et al., 2009; Pappas et al., 2010; Ochala et al., 2012). Ongoing work seeks to understand the regulatory framework governing the interplay between aging and pathology of skeletal muscle, alterations in thin filament proteins, and regulation of thin filament lengths.

#### **ACKNOWLEDGMENTS**

This work was supported by stipends from the Summer Undergraduate Research Fellowship program at The Scripps Research Institute (to Emily A. Dubuc and Kendra Q. Lian), a development grant from the Muscular Dystrophy Association (to David S. Gokhin), an NIH/NIAMS Pathway to Independence award (K99-AR066534 to David S. Gokhin), a grant from NIH/NIA to the Nathan Shock Center of Excellence in the Basic Biology of Aging at The Jackson Laboratory (P30-AG038070 to Luanne L. Peters), and a grant from NIH/NIAMS to the Imaging Core of the San Diego Skeletal Muscle Research Center (P30-AR061303 to Velia M. Fowler).

#### **REFERENCES**

indirect flight muscle. *J. Cell Biol.* 155, 1043–1053. doi: 10.1083/jcb. 200108026

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 27 August 2014; accepted: 10 September 2014; published online: 29 September 2014.*

*Citation: Gokhin DS, Dubuc EA, Lian KQ, Peters LL and Fowler VM (2014) Alterations in thin filament length during postnatal skeletal muscle development and aging in mice. Front. Physiol. 5:375. doi: 10.3389/fphys.2014.00375*

*This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology.*

*Copyright © 2014 Gokhin, Dubuc, Lian, Peters and Fowler. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## O-GlcNAcylation, contractile protein modifications and calcium affinity in skeletal muscle

#### *Caroline Cieniewski-Bernard1,2, Matthias Lambert 1,2, Erwan Dupont 1,2, Valérie Montel 1,2, Laurence Stevens 1,2 and Bruno Bastide1,2\**

*1 Université Lille, Lille, France*

*2 EA4488, APMS, URePsss, Université de Lille 1, Villeneuve d'Ascq, France*

#### *Edited by:*

*Julien Ochala, King's College London, UK*

#### *Reviewed by:*

*Christina Karatzaferi, University of Thessaly, Greece Ranganath Mamidi, Case Western Reserve University, USA*

#### *\*Correspondence:*

*Bruno Bastide, EA4488 Laboratoire Activité Physique, Muscle et Santé, Biology Department, Université de Lille 1, Bât SN4, UFR Biology, 59655 Villeneuve d'Ascq, France e-mail<EMAIL_ADDRESS>
O-GlcNAcylation, a generally undermined atypical protein glycosylation process, is involved in a dynamic and highly regulated interplay with phosphorylation. Akin to phosphorylation, O-GlcNAcylation is also involved in the physiopathology of several acquired diseases, such as muscle insulin resistance or muscle atrophy. Recent data underline that the interplay between phosphorylation and O-GlcNAcylation acts as a modulator of skeletal muscle contractile activity. In particular, the O-GlcNAcylation level of the phosphoprotein myosin light chain 2 seems to be crucial in the modulation of the calcium activation properties, and should be responsible for changes in calcium properties observed in functional atrophy. Moreover, since several key structural proteins are O-GlcNAc-modified, and because of the localization of the enzymes involved in the O-GlcNAcylation/de-O-GlcNAcylation process to the nodal Z disk, a role of O-GlcNAcylation in the modulation of the sarcomeric structure should be considered.

**Keywords: O-GlcNAcylation, phosphorylation, O-GlcNAcylation/phosphorylation interplay, contractile proteins, MLC2, contractile properties, sarcomeric structure**

### **O-GlcNAcylation, AN ATYPICAL GLYCOSYLATION**

Nowadays, it is well admitted that the phosphorylation does not act alone in the fine modulation of numerous cellular processes, but rather, presents a dynamic and highly regulated interplay with an atypical glycosylation, the O-linked Nacetyl-glucosaminylation (termed O-GlcNAcylation), occurring on nuclear, cytoplasmic and mitochondrial proteins (Hart et al., 2007; Cao et al., 2013; Johnsen et al., 2013). This minireview is based on significant references focused on the recent advancements concerning the link between O-GlcNAcylation, contractile proteins and calcium affinity in skeletal muscle (**Figure 1**).

The O-GlcNAcylation of proteins results from the transfer of N-acetyl-β-D-glucosamine from the high energy donor substrate UDP-GlcNAc (synthesized through the hexosamine biosynthesis pathway) onto the hydroxyl group of serine and threonine residues of target proteins by the uridine diphospho-Nacetyl glucosaminyl transferase (O-GlcNAc transferase or OGT). The β-N-acetylglucosaminidase (O-GlcNAcase or OGA) catalyzes the removal of O-GlcNAc residues from proteins (Dong and Hart, 1994; Gao et al., 2001; Wells et al., 2002). Thus, like phosphorylation, the addition/removal of GlcNAc moieties on the proteins results from the concerted action of two antagonist enzymes. Reversible, O-GlcNAcylation is highly dynamic, and responds rapidly to changes in environmental conditions (Hart et al., 2007). Since OGT is coded from only one gene (Kreppel et al., 1997), the regulation of its activity, its localization in cell, or its specificity toward protein targets are ensured by targeting proteins, such as protein phosphatase 1 (PP1), Milton (OIP106), p38MAP kinase, myosin phosphatase 1 (MYPT1) or peroxisome-proliferator-activated receptor-co-activator-1alpha (PGC1α), transiently associated with OGT through tetratricopeptide repeats (TPR domains) at the N-termini of the transferase (Iyer and Hart, 2003; Iyer et al., 2003; Wells et al., 2004; Cheung and Hart, 2008; Cheung et al., 2008; Housley et al., 2009). In the same way, the OGA could also be associated with other proteins like calcineurin or heat shock proteins among others (Wells et al., 2001). Moreover, an unusual association of the two opposing OGA/OGT was described, forming a single O-GlcNAczyme complex (Whisenhunt et al., 2006).

The analysis of O-GlcNAc pattern, the quantification of variation of O-GlcNAcylation on proteins and the identification of the glycosylated sites are crucial for the understanding of the role of this atypical glycosylation. Methodological approaches include western blot analyses using antibodies directed against O-GlcNAc moieties or lectins (Zachara et al., 2011), or the labeling of O-GlcNAcylated proteins with galactosyltransferase and coupling of different kind of substrates (Zachara et al., 2011). Moreover, the identification and mapping of O-GlcNAc modification sites have been at the origin of several technical developments during the 10 last years (Ma and Hart, 2014 for review).

O-GlcNAcylation has been shown to be involved in almost all cellular processes, including signal transduction, protein degradation or regulation of gene expression (Hart et al., 2011; Bond and Hanover, 2013). O-GlcNAcylation has been also demonstrated to act as an inducible, cytoprotective stress response. Indeed, increase in O-GlcNAcylation of nucleocytoplasmic proteins

protects cells with an induction of heat shock proteins (Zachara et al., 2004). In cardiomyocytes, O-GlcNAcylation can attenuate oxidative stress through inhibition of calcium overload and ROS generation (Ngoh et al., 2010). Studies performed with OGT and OGA knockout mice have demonstrated that O-GlcNAc is crucial for life since O-GlcNAcylation was essential for embryonic stem cell viability and is implicated in the aging process, elevation of O-GlcNAcylation being altered in different tissues of different ages (Shafi et al., 2000; Yang et al., 2012). Moreover, many reports demonstrate that O-GlcNAcylation might play a role in the physiopathology of several acquired diseases, such as cancer, cardiovascular diseases, neurodegenerative diseases, Alzheimer or type II diabetes (Lefebvre et al., 2010; Slawson et al., 2010; Nakamura et al., 2012; Bond and Hanover, 2013; Ma and Hart, 2013; Ma and Vosseller, 2013; Förster et al., 2014). Indeed, increased O-GlcNAcylation is closely linked to insulin resistance and hyperglycemia-induced glucose toxicity.

#### **O-GlcNAcylation, SKELETAL MUSCLE, AND INSULIN-RESISTANCE**

The concept of a role of O-GlcNAcylation in skeletal muscle has emerged from studies considering muscle as one of the crucial insulin-sensitive tissue; indeed, skeletal muscle is responsible for more than 80% of insulin-stimulated glucose uptake in humans. In the past decades, studies in rodents (Hawkins et al., 1997a,b) suggested a correlation between the development of insulin resistance and increased UDP-GlcNAc concentrations in muscle. Thus, raising O-GlcNAc level in skeletal muscle has been demonstrated to induce insulin resistance (Arias and Cartee, 2005, **Figure 1**), while coinfusion of insulin and glucosamine, increasing UDP-GlcNAc, enhances the O-GlcNAc modification on numerous unidentified skeletal muscle proteins (Yki-Jarvinen et al., 1998). Moreover, transgenic mice overexpressing GLUT1 in skeletal muscle was insulin resistant and exhibit chronically increased glucose flow and increased UDP-GlcNAc concentrations in muscle (Buse et al., 1996); indeed, 2–5% of the glucose entering into the cell is directed to the hexosamine biosynthesis pathway, leading to the synthesis of UDP-GlcNAc, the donor for O-GlcNAcylation. More recently, it has been suggested that mitochondrial and contractile dysfunctions, observed in the development of Type 2 diabetes, were linked to the increase in O-GlcNAcylation level (Johnsen et al., 2013). Thus, in rats artificially selected for Low Running Capacity (LCR rats), predisposed to becoming obese, and developing insulin resistance and cardiovascular dysfunction, the increase in O-GlcNAcylation on mitochondrial proteins and SERCA was associated with mitochondrial dysfunction and changes in contractile properties (Koch et al., 2012). In the LCR heart myocytes, the cardiovascular dysfunction could be attributed to the decline in the contractility, correlated to the impaired intracellular Ca2+ handling and signaling (Koch et al., 2012). Hu et al. (2005) demonstrated also the correlation between O-GlcNAcylation and diabetes since they demonstrated that adenoviral transfer of OGA into the myocardium of streptozotocin induced diabetic mice, reversed the excessive O-GlcNAc modifications associated with diabetes, particularly in the contractile dysfunctions and Ca2+ handling capacities (Hu et al., 2005). The decline in contractibility could be related to the decrease in calcium affinity and sensitivity reported in skinned cardiac fibers in presence of GlcNAc (Ramirez-Correa et al., 2008) but rather involved changes in expression of SERCA (Hu et al., 2005; Johnsen et al., 2013). This role of O-GlcNAcylation should be considered also in skeletal muscle since dysfunction of contractibility as well as the Ca2+ handling were also measured in skeletal muscle of rat models of diabetes (Eshima et al., 2013), via impaired SERCA and Glut 4 (Safwat et al., 2013).

#### **O-GlcNAcylation/PHOSPHORYLATION INTERPLAY AND SKELETAL MUSCLE CONTRACTILE ACTIVITY**

Phosphorylation was well-admitted for a long time to regulate many key processes in muscle physiology. Among them, phosphorylation is a key regulator of the intracellular signaling pathways involved in muscle mass and phenotype adaptation to physiological demands. Moreover, phosphorylation is involved in myofibrillar physiology, such as muscular contraction or cellular structuration through the modulation of protein-protein interactions. Indeed, phosphorylation has been demonstrated to play a crucial role in the regulation of the contractile properties in striated muscle, but recent reports suggested that O-GlcNAcylation might play a role as important as phosphorylation in muscle physiology. Thus, many proteins of muscle proteome have been identified to be O-GlcNAc modified (Cieniewski-Bernard et al., 2004, 2012; Hedou et al., 2007; Ramirez-Correa et al., 2008). Among them, several key contractile proteins of striated muscle are concerned, i.e., myosin heavy chains (slow MHCI as well as the fast isoforms MHCIIA and MHCIIB), myosin light chains (essential MLC or MLC1 and regulatory MLC or MLC2), actin, both α and β isoforms of tropomyosin as well as isoforms of TnI and TnT. It is noteworthy that it is not known whether TnC, the calcium-sensor, bears an O-GlcNAc moiety or not.

While contraction is triggered by calcium release, it is regulated by several myofilament proteins such as the regulatory myosin light chain (termed MLC2). MLC2 phosphorylation is not essential for skeletal muscle contraction but is an important regulatory mechanism since it can produce changes in thick filament structure and enhance crossbridge attachment (Szczesna et al., 2002). For instance, phosphorylation of MLC2, catalyzed by a Ca2+/calmodulin-dependant MLC kinase, increases the force development at submaximal calcium concentration, conferring a higher calcium sensitivity to the fibers (Persechini et al., 1985; Stephenson and Stephenson, 1993; Sweeney et al., 1993; Szczesna et al., 2002).

Our recent data highlight the key role of O-GlcNAcylation as a modulator of skeletal muscle contractile activity, in particular on the calcium activation properties (Hedou et al., 2007; Cieniewski-Bernard et al., 2012). Indeed, muscle skinned fibers, when exposed to N-acetyl-D-glucosamine, present a reversible decrease in calcium sensitivity and affinity, whereas the cooperativity within the thin filament was not changed (Hedou et al., 2007). This modulation probably involved disruption of proteinprotein interactions through O-GlcNAc moieties. Interestingly, a similar effect was observed in rats as well as in human skinned fibers (Cieniewski-Bernard et al., 2009) and also measured in cardiac trabeculae (Ramirez-Correa et al., 2008).

Further experiments performed after the pharmacological increase of O-GlcNAcylation level of contractile protein glycosylation, using PUGNAc or Thiamet G, two inhibitors of OGA (Gloster and Vocadlo, 2010), leads to an increase of calcium affinity on slow soleus skinned fibers (Cieniewski-Bernard et al., 2012). Several regulatory contractile proteins, predominantly fast isoforms, presented a drastic increase in their O-GlcNAc level. Since the only slow isoform of contractile protein (and so the more representative isoforms in the slow skeletal muscle) presenting an increase of O-GlcNAc level was the myosin regulatory light chains MLC2, the effect of enhanced O-GlcNAcylation pattern on calcium activation parameters of slow soleus fibers was attributed to the increase of the O-GlcNAcylation of sMLC2 (Cieniewski-Bernard et al., 2012).

All these data closely linked O-GlcNAcylation to the modulation of contractile activity of skeletal muscle, a decrease in O-GlcNAcylation being associated to a decrease in calcium affinity and reciprocally (**Figure 1**). Moreover, analysis of the proteins presenting changes in their O-GlcNAc level states suggests that MLC2 could play an important role in this modulation. Since MLC2 was identified to be O-GlcNAcylated (Hedou et al., 2007), and because of the potential antagonism between phosphorylation and O-GlcNAcylation, i.e., the potential O-GlcNAcylated site corresponds to the only site of phosphorylation in MLC2 located in Ser 14 for the rat slow MLC2 isoform (Blumenthal and Stull, 1980; Ramirez-Correa et al., 2008), the O-GlcNAcylation should be considered as a potential new mechanism that could modulate the contractile properties of skeletal muscle as well as the phosphorylated states of MLC2.

#### **O-GlcNAcylation/PHOSPHORYLATION INTERPLAY, AND SKELETAL MUSCLE CONTRACTILE DYSFUNCTION IN ATROPHIED MUSCLE**

Muscle atrophy characterized skeletal muscle adaptation to a large variety of disuse conditions (immobilization, microgravity, bed rest, or nerve injury). This atrophy results from a reduction in fiber diameter, protein content and is associated with slow to fast phenotype transitions accompanied by functional changes such as loss in force and increased fatigability (Baldwin et al., 1996; Fitts et al., 2000; Fluck and Hoppeler, 2003; Mounier et al., 2009). Moreover a role of O-GlcNAcylation in the development of muscle atrophy has been suggested (Cieniewski-Bernard et al., 2006; **Figure 1**). Indeed, a correlation between variations in O-GlcNAcylation levels and the development of atrophy after hindlimb unloading was shown, suggesting that O-GlcNAc variations could control the muscle protein homeostasis (Cieniewski-Bernard et al., 2006). In particular, it was suggested that O-GlcNAcylation should be implicated in the regulation of muscular atrophy as a protective mechanism against proteasomal degradation (Cieniewski-Bernard et al., 2006). It was also recently described that muscle-specific overexpression of NCOATGK, a splice variant of O-GlcNAcase, induces skeletal muscle atrophy (Huang et al., 2011).

We have demonstrated that the slow-to-fast transition (at MHC and MLC2 level) concomittantly to soleus muscle atrophy was correlated with an increase of the global level of MLC2 phosphorylation in rat hindlimb unloading model (Bozzo et al., 2005) as well as in human patients who were subjected to 2-months Bed Rest (Stevens et al., 2013). However, similar variations in MLC2 phosphorylation have been observed when clenbuterol was administrated to rats; in such case, muscle hypertrophy was associated to slow-to-fast transition in rats (Bozzo et al., 2003).

Thus, the slow-to-fast MLC2 transition was associated to an increase of phosphorylation level of the two isoforms regardless of whether hypertrophy or atrophy develops. Moreover, although numerous data argue that MLC2 phosphorylation lead to higher calcium sensitivity in skeletal muscle, an increase in MLC2 phosphorylation was measured after muscle atrophy whereas the calcium sensitivity decreases (Bozzo et al., 2003). Similar data were obtained from phosphoproteome analysis in human aged muscle (Gannon et al., 2008). These data could indicate that the decrease in calcium sensitivity occurring during hindlimb unloading could involve another type of regulation than classical phosphorylation. Since we demonstrated that a decrease in calcium affinity was measured when O-GlcNAcylation is reduced, and since a decrease in O-GlcNAcylation is observed in atrophied muscle, one hypothesis is that O-GlcNAcylation might be involved in this alteration.

To support this hypothesis, the variation of O-GlcNAcylation level on MLC2, as well as its interplay with phosphorylation, was investigated in atrophied soleus muscle (Cieniewski-Bernard et al., in press). Interestingly, an antagonism between phosphorylation and O-GlcNAcylation was demonstrated on the slow MLC2 in a rat model of hindlimb unloading, largely used to induce functional atrophy of antigravitary muscles such as soleus (Cieniewski-Bernard et al., in press), since a decrease in MLC2 O-GlcNAcylation was measured in atrophied soleus associated to the increase in phosphorylation states as previously described. More importantly, the use of Phos-Tag acrylamide gels allowed the authors to analyze the O-GlcNAcylation state on each phosphorylated form of MLC2 (Cieniewski-Bernard et al., in press). It was demonstrated that the two post-translational modifications were mutually exclusive, and that this interplay was closely associated to load because of the reversibility of these processes occurring during reloading. Interestingly, it was demonstrated that the enzymes involved in the phosphorylation/dephosphorylation and O-GlcNAcylation/de-O-GlcNAcylation processes were associated within a multi-enzymatic complex, and were localized to the nodal Z disk region of the sarcomere. All these results are in favor of an interplay between phosphorylation and O-GlcNAcylation on MLC2, probably through the same site.

This interplay between phosphorylation and O-GlcNAcylation in the fine modulation of MLC2 activity was also observed in humans (Stevens et al., 2013). Indeed, the post-translational modifications of MLC2 were investigated in soleus biopsies obtained from 60-days Bed-Rest female subjects. In this cohort, several groups were formed: 60-days Bed Rest (BR), BR + Exercice (combined aerobic and resistive exercises), and BR + Nutritional protocol (leucine and valine diet). The slow-fast phenotype transition of MLC2 was associated to an increase of the phosphorylation states of slow and fast isoforms MLC2 while the global MLC2 glycosylation level was decreased. Interestingly, aerobic and resistive exercises, which preserved muscles from BR changes (and so slow-to-fast transition), also prevented this antagonism.

Taken together, all these data clearly corroborated the interplay between phosphorylation and O-GlcNAcylation on MLC2, in animal and in human models of functional atrophy. Indeed, the decrease in calcium sensitivity as well as calcium affinity measured in skinned fibers of atrophied muscle soleus cannot be obviously explained by the increase in MLC2 phosphorylation state, as it was previously described (Bozzo et al., 2003). Since previous experiments have demonstrated that decrease in O-GlcNAcylation was associated to a decrease in calcium sensitivity and affinity, whereas its phosphorylated state increased in atrophied muscle, MLC2 O-GlcNAcylation state could be predominantly involved in the alteration of contractile parameters observed in atrophied soleus. In contrast, the MLC2 phosphorylation might be rather associated to phenotype transition than to changes in contractile parameters in atrophied muscle. Interestingly, some data support for a structural role of MLC2 phosphorylation: MLC2 phosphorylation induces stress fiber assembly in nonmuscle cells (Katoh et al., 2001) and mediates sarcomere organization during hypertrophic growth in cardiac muscle (Aoki et al., 2000).

However, further functional experiments need to be performed to demonstrate unambiguously that the phosphorylation/O-GlcNAcylation balance of MLC2 might be directly involved in the decrease in calcium sensitivity and affinity observed in atrophied muscles. While recent reports focused on MLC2, other regulatory contractile proteins have been demonstrated to be O-GlcNAcylated such as Tropomyosin, TnT and TnI (Cieniewski-Bernard et al., 2012). We cannot totally exclude that the glycosylation state of these proteins nor that unidentified O-GlcNAc proteins (more particularly, TnC is one of them) might play a role in the modulation of contractile activity.

#### **MYOFIBRILLAR PROTEINS O-GlcNAcylation AND PERSPECTIVES**

From the data described above, it seems very likely that O-GlcNAcylation plays a major role in modulating the contractile activity in skeletal muscle. However, the role of the O-GlcNAcylation might also concern other mechanisms involved in the sarcomeric structure. The preferential localization of OGT and OGA in the sarcomere, more particularly at the Z disk region (Cieniewski-Bernard et al., in press), argue for an important role of O-GlcNAcylation in this nodal region that could influence mechanisms other than the modulation of the contractile activity. Indeed, it was demonstrated that myosin, actin but also key proteins involved in the sarcomeric structure (desmin, actinin, αB-crystallin, and ZASP) were modified by O-GlcNAc moieties (Cieniewski-Bernard et al., 2012; Leung et al., 2013). Interestingly, we demonstrated that the O-GlcNAc sites on myosin were localized in a region involved in polymerization and interaction of myosin with proteins partner such as myomesin, M-protein or titin (Hedou et al., 2009) supporting a role for O-GlcNAcylation in the organization of the sarcomere (**Figure 1**). This hypothesis is underlined by the fact that numerous structural proteins are modified by O-GlcNAc: beta3 integrin (Ahmad et al., 2006), vinculin (Laczy et al., 2010), spectrin (Zhang and Bennett, 1996), alphaB-crystallin (Roquemore et al., 1996), laminin (Kwak et al., 2010), or cytokeratin (Ku and Omary, 1995), and variation of O-GlcNAcylation of the intermediate filaments has been demonstrated to lead to defective phosphorylated states and so to polymerization (Farah and Galileo, 2008; Slawson et al., 2008). Moreover, O-GlcNAcylation modulates organization and solubilization of cytokeratin (Rotty et al., 2010; Srikanth et al., 2010). Interestingly, It has been demonstrated that the intermediate filament proteins, vimentin and desmin, possess lectin-like properties toward O-GlcNAc moieties, supporting physiological interaction between GlcNAc-bearing ligands (and so O-GlcNAc proteins) and lectinic proteins (Ise et al., 2010). The assembly and regular arrangement of the sarcomere results from highly regulated interactions between myofibrillar proteins and structural proteins which are at the origin of a sarcomeric cytoskeleton. It's becoming evident from recent data that a role of O-GlcNAcylation in the organization of the sarcomere should be considered. Since phosphorylation has been demonstrated to be involved in protein-protein interactions, a balance between phosphorylation and O-GlcNAcylation might modulate the dynamics of the sarcomere structural organization. Future investigations will be conducted with aim to determine the role of O-GlcNAcylation in the organization and dynamic of the sarcomere. This work may provide new insights in the understanding of molecular mechanisms of diseases characterized by a disintegration of myofibrils and marked disorganization of the Z band region such as myofibrillar and congenital myopathies.

#### **ACKNOWLEDGMENTS**

This work was supported by grants from the Région Nord-Pasde-Calais 2011 (Emergent Research Project n◦12003808) and the CNES (French Spatial Agency, n◦4800000716).

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O-GlcNAcylation blocks signaling and development. *Glycobiology* 16, 551–563. doi: 10.1093/glycob/cwj096

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 08 September 2014; accepted: 11 October 2014; published online: 30 October 2014.*

*Citation: Cieniewski-Bernard C, Lambert M, Dupont E, Montel V, Stevens L and Bastide B (2014) O-GlcNAcylation, contractile protein modifications and calcium affinity in skeletal muscle. Front. Physiol. 5:421. doi: 10.3389/fphys.2014.00421*

*This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology.*

*Copyright © 2014 Cieniewski-Bernard, Lambert, Dupont, Montel, Stevens and Bastide. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Pseudo-acetylation of K326 and K328 of actin disrupts Drosophila melanogaster indirect flight muscle structure and performance

Meera C. Viswanathan, Anna C. Blice-Baum, William Schmidt, D. Brian Foster and Anthony Cammarato\*

Division of Cardiology, Department of Medicine, Johns Hopkins University School of Medicine, Baltimore, MD, USA

#### Edited by:

Julien Ochala, King's College London, UK

#### Reviewed by:

Frieder Schoeck, McGill University, Canada Richard Cripps, University of New Mexico, USA

#### \*Correspondence:

Anthony Cammarato, Division of Cardiology, Department of Medicine, Johns Hopkins University School of Medicine, 720 Rutland Avenue, Ross 1050, Baltimore, MD 21205, USA<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Striated Muscle Physiology, a section of the journal Frontiers in Physiology

Received: 07 November 2014 Accepted: 26 March 2015 Published: 28 April 2015

#### Citation:

Viswanathan MC, Blice-Baum AC, Schmidt W, Foster DB and Cammarato A (2015) Pseudo-acetylation of K326 and K328 of actin disrupts Drosophila melanogaster indirect flight muscle structure and performance. Front. Physiol. 6:116. doi: 10.3389/fphys.2015.00116 In striated muscle tropomyosin (Tm) extends along the length of F-actin-containing thin filaments. Its location governs access of myosin binding sites on actin and, hence, force production. Intermolecular electrostatic associations are believed to mediate critical interactions between the proteins. For example, actin residues K326, K328, and R147 were predicted to establish contacts with E181 of Tm. Moreover, K328 also potentially forms direct interactions with E286 of myosin when the motor is strongly bound. Recently, LC-MS/MS analysis of the cardiac acetyl-lysine proteome revealed K326 and K328 of actin were acetylated, a post-translational modification (PTM) that masks the residues' inherent positive charges. Here, we tested the hypothesis that by removing the vital actin charges at residues 326 and 328, the PTM would perturb Tm positioning and/or strong myosin binding as manifested by altered skeletal muscle function and structure in the Drosophila melanogaster model system. Transgenic flies were created that permit tissue-specific expression of K326Q, K328Q, or K326Q/K328Q acetyl-mimetic actin and of wild-type actin via the UAS-GAL4 bipartite expression system. Compared to wild-type actin, muscle-restricted expression of mutant actin had a dose-dependent effect on flight ability. Moreover, excessive K328Q and K326Q/K328Q actin overexpression induced indirect flight muscle degeneration, a phenotype consistent with hypercontraction observed in other Drosophila myofibrillar mutants. Based on F-actin-Tm and F-actin-Tm-myosin models and on our physiological data, we conclude that acetylating K326 and K328 of actin alters electrostatic associations with Tm and/or myosin and thereby augments contractile properties. Our findings highlight the utility of Drosophila as a model that permits efficient targeted design and assessment of molecular and tissue-specific responses to muscle protein modifications, in vivo.

Keywords: tropomyosin, myosin, acetylation, muscle contraction, post-translational modification

## Introduction

Striated muscle contraction results from transient interactions between myosin-containing thick filaments and actin-containing thin filaments. Contractile regulation is achieved by Ca2+ dependent modulation of myosin S1 cross-bridge binding to actin by the thin filament-associated troponin-tropomyosin complex (reviewed in Tobacman, 1996; Gordon et al., 2000; Brown and Cohen, 2005; Lehman and Craig, 2008). The location of continuous troponin-tropomyosin complexes along the surface of F-actin governs the access of myosin binding sites and, hence, force production (Haselgrove, 1973; Huxley, 1973; Parry and Squire, 1973; McKillop and Geeves, 1993; Lehman et al., 1994; Vibert et al., 1997). Under conditions of low Ca2+, the troponin complex constrains tropomyosin (Tm) in a position that occludes myosin target sites on actin. Consequently, Tm sterically blocks and limits myosin binding, and relaxation results. During muscle activation, Ca2+ binds to troponin and triggers movement of Tm away from myosin binding sites. This relocation partially relieves the structural blocking imposed by Tm. Initial myosin binding on thin filaments further displaces Tm and exposes myosin binding sites along F-actin, thereby contributing to the cooperative activation of contraction.

Although the specific residues that constitute the binding interface of actin and Tm are not completely known, it is well accepted that the association of Tm with actin is largely electrostatic (Lorenz et al., 1995; Brown and Cohen, 2005; Barua et al., 2011, 2013; Li et al., 2011). Recently, models of the conserved binding interface between actin and Tm in various states have been proposed based on molecular evolutionary and mutational analysis, computational chemistry, and electron microscopy reconstructions (Barua et al., 2011, 2013; Li et al., 2011; Behrmann et al., 2012; Von Der Ecken et al., 2015). Structural studies of F-actin-Tm and F-actin-Tm-myosin revealed that several amino acids on actin can potentially form distinct contacts with Tm in the absence and presence of myosin S1 (Li et al., 2011; Behrmann et al., 2012; Von Der Ecken et al., 2015). For example, in the absence of S1, a cluster of basic actin residues comprised of K326, K328, and R147 appeared poised to clasp onto E181 of Tm to establish highly favorable associations (**Figure 1**) (Li et al., 2011). Interestingly, K328 also interacts electrostatically with E286 of S1 to help define a strong contact point between actin and rigor-bound myosin (Behrmann et al., 2012).

Alterations to thin filament proteins affect the properties of muscle. Missense mutations, for instance, can disrupt conserved interfaces among cardiac thin filament subunits and initiate diverse cardiomyopathies (Tardiff, 2011). Similar to diseasecausing mutations, post translational modifications (PTMs) also alter the chemical nature of thin filament subunits. Although less-well appreciated, these modifications are widely employed in vivo, occur through enzymatic and non-enzymatic mechanisms, and direct both physiological and pathological processes (Terman and Kashina, 2013). PTMs add or remove a functional group to or from specific amino acid residues, which can induce changes in protein structure, activity, or binding partners (van Eyk, 2011). To date, more than 400 different PTMs have been described, although far fewer have been documented in higher organisms (Agnetti et al., 2011). In the cardiac subproteome, phosphorylation is by far the best-described PTM (Sumandea et al., 2004; Agnetti et al., 2011; Solaro and Kobayashi, 2011; van Eyk, 2011). It has been observed for 80% of the myofilamentous proteins (Agnetti et al., 2011). However, the effects of the majority of the potential PTMs have not been fully

FIGURE 1 | Critical electrostatic F-actin-Tm and F-actin-Tm-myosin interactions. (A) Molecular models showing the location of tropomyosin (Tm) (blue) on actin (yellow and green) in the absence and presence of the myosin head (S1) (red) bound in rigor. The F-actin-Tm and rigor F-actin-Tm-myosin structures are based on those generated by Li et al. (2011) and Behrmann et al. (2012) respectively. (B) Enlarged views illustrate critical electrostatic associations between actin and Tm in the absence or

presence of S1. K328 on actin (circled) interacts with E181 of Tm in the absence of myosin (left) and with E286 of myosin when S1 is bound in rigor (right). Note the azimuthal movement of Tm across F-actin. (C) Projected and enlarged views highlight vital electrostatic interactions of actin residues R147, K326, and K328 with E181 of Tm, in the absence of myosin, and of K328 of actin and E286 of S1 when myosin is bound in rigor. These associations are likely critical for thin filament and muscle function.

investigated, in part because of a lack of technologies needed to target and reliably identify them. Moreover, compared to inherited mutations, the influence of relatively few PTMs on contractile performance have been examined in the physiological context of muscle.

Actin is an abundant and highly conserved protein (**Figure 2**). It participates in more protein-protein interactions than any known protein and is subject to a number of PTMs (Herman, 1993; Dominguez and Holmes, 2011; Terman and Kashina, 2013). Characterizing these modifications constitutes a rapidly expanding area within actin studies and muscle biology (Terman and Kashina, 2013). Phosphorylation, methylation, ADP-ribosylation, oxidation, arginylation, O-GlcNAcylation, ubiquitylation, and acetylation are examples of major actin modifications, many of which have now been confirmed in sarcomeric actin. For example, K326 and K328 of actin isolated from various cell lines were shown to be acetylated (Choudhary et al., 2009) and, as recently described by Foster et al. (2013) these residues were also shown to be acetylated on actin recovered from the myofilament fraction of guinea pig hearts. Acetylation is a reversible PTM that neutralizes the positive charge of K326 and K328 on actin. Unlike the N-terminal acetylation, acetylation of these residues has unknown function. Based on in vitro and in silico structural predictions, removal of positively charged amino acids that are potentially critical to Tm and myosin electrostatic associations likely alters muscle performance substantially and in diverse ways.

Here, we aimed to investigate the in vivo physiological and morphological consequences of acetylating actin residues K326 and K328 on muscle. We chose the indirect flight muscle (IFM) of D. melanogaster as our primary experimental vehicle. Drosophila IFMs are highly amenable to mechanical and structural analyses, are not required for viability, exhibit a stretch activation response that is similar to that of cardiac muscle, and provide ample material that is easily isolated for biochemical and biophysical experimentation (Bing et al., 1998; Razzaq et al., 1999; Cammarato et al., 2004; Vikhorev et al., 2010; Swank, 2012). The vast array of genetic tools available to manipulate the fly's genome also provides unique opportunities to examine how thin filament modifications affect muscle structure and performance. The GAL4-UAS system simplifies tissue-specific expression of mutant transgenes (Brand and Perrimon, 1993). However, previous studies suggested that IFM myofibril assembly and flight ability are highly sensitive to the stoichiometry of muscle proteins (Beall and Fyrberg, 1991; Bernstein et al., 1993), and therefore, the utility of this bipartite expression system for investigating major contractile components has been questioned. For example, despite normal sarcomere appearance, overexpression of several UAS-GFP-actin alleles was shown to exclusively affect IFM function, leading to flightless adults that were otherwise healthy (Röper et al., 2005). The effects of non-GFP fusion constructs on IFM performance were not tested. Therefore, to determine if the IFM can serve as a viable, conditional model system for probing the effects of PTMs on myofibrillar components, we tested the hypotheses that (1) GAL4-UAS-mediated overexpression of a Drosophila cardiac actin isoform in the IFM would not perturb gross morphology or flight performance as determined by standard metrics and that (2) expression of K326Q, K328Q, or K326Q/K328Q acetyl-mimetic cardiac actin disrupts vital electrostatic interactions required for Tm positioning and/or strong myosin binding, in vivo, as manifested by perturbed IFM structure and function. Using four muscle-specific GAL4 drivers we found that expressing wild-type Drosophila cardiac actin had no effect on flight or gross muscle organization. Relatively high expression levels of K326Q cardiac actin mildly affected flight ability, whereas excessive amounts of K328Q and K326Q/K328Q cardiac actin eliminated flight and triggered IFM destruction. Based on F-actin-Tm and F-actin-Tm-myosin models and on our physiological data, we propose that acetylating K326 and K328 of actin alters crucial electrostatic associations with Tm and/or myosin and thereby promotes actomyosin associations and modulates muscle performance. Our findings highlight the utility of Drosophila as a model that permits efficient targeted design and assessment of tissue-specific responses to muscle protein modifications, in vivo.

## Materials and Methods

#### Structural Modeling

Models of the F-actin-Tm (Li et al., 2011; Orzechowski et al., 2014) and the rigor F-actin-Tm-myosin (PDB ID: 4A7F) (Behrmann et al., 2012) binding interfaces were generated using the molecular modeling program, Chimera version 1.9 (Pettersen et al., 2004).

#### Multiple Sequence Alignment

Sequence comparison of skeletal and cardiac actin isoforms from Homo sapiens, Cavia porcellus, and D. melanogaster was performed using the Clustal Omega multiple sequence alignment program. Residues were shaded based on degree of conservation.

#### Fly Stocks

All flies were raised at 25◦C on a standard cornmeal-yeastsucrose-agar medium. The w 1118 strain was obtained from Genetic Services Inc. (Sudbury, MA) and the Mef2-GAL4 driver line (y 1 w ∗ ; P{GAL4-Mef2.R}3) from the Bloomington Drosophila Stock Center (Bloomington, IN). The MHC-GAL4 (MHC-GAL482) line described by Marek et al. (2000) was obtained from Dr. Rolf Bodmer (Sanford Burnham Medical Research Institute, La, Jolla, CA). The UH3-GAL4 (Singh et al., 2014) and the Act88F-GAL4 (88F2) (Bryantsev et al., 2012) lines were gifts from Dr. Upendra Nongthomba (Indian Institute of Science, Banglore, India) and Dr. Richard M. Cripps (University of New Mexico, Albuquerque, NM) respectively. The Mhc10 (w; Mhc10/Mhc10; TM2/MKRS) IFM-specific myosin null line was acquired from Dr. Sanford I. Bernstein (San Diego State University, San Diego, CA). Mef2-GAL4> UAS-Act57BWT; Mhc10/+ and UAS-Act57BK328Q; Mhc10/+ Drosophila were generated by standard mating procedures.

#### Construction of UAS-Actin Transgenes

The N-terminally-labeled GFP-actin construct (pUASp.Act57BGFP.WT) was generously provided by Dr. Katja Röper (MRC- Laboratory of Molecular Biology, Cambridge, UK). The Act57BGFP.WT and Act57BWT cDNA sequences were subsequently inserted into the pUASTattB vector (obtained from Dr. Christopher Potter, Johns Hopkins University) using the KpnI and XbaI and the NotI and XbaI restriction sites respectively. The pUASTattB vector includes the Drosophila miniwhite (w +) gene as a selectable eye color marker. The Act57B actin acetyl-mimetic mutations, K326Q, K328Q, and K326Q/K328Q, were generated by site-directed mutagenesis using specific primer pairs and the QuikChange Site-directed mutagenesis kit (Agilent Technologies).

List of primers for site-directed mutagenesis:

#### Generation of Transgenic Drosophila

The pUASTattB constructs were injected into attp40 Drosophila embryos for PhiC31 integrase mediated site-specific transgenesis (transgene landing site cytolocation 25C7) by Genetic Services, Inc. Injected adults were crossed to w 1118 flies and the progeny screened for pigmented eye color, which reflects the presence of the miniwhite (w +) marker and the transgene, an indicator of successful transformation. Each transformant fly was then crossed into the w 1118 background to generate stable transgenic lines.

## Verification of Transgene Expression

Transgenic actin expression was verified by isolating total RNA from bisected thoraces or dissected IFMs of 10 Mef2-GAL4> UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, or UAS-Act57BK326Q/K328Q flies using the Quick-RNA microprep kit (Zymo Research Corp). Contaminating DNA was removed with RNase free DNase I (Zymo Research Corp). One step RT-PCR was carried out with the Qiagen QuantiTect Reverse Transcription Kit (Qiagen Inc) and 10 ng RNA per reaction. The cDNA was amplified using an Act57B primer pair (5′ CCCTGTACGCCTCCGGTCGTA 3′ and 5′ TTAGAAGCACTTGCGGTGGAC 3′ ) and the amplified product was sequenced at the Johns Hopkins Synthesis and Sequencing facility.

Transgenic muscle-restricted protein expression was confirmed using the UAS-Act57BGFP.WT reporter line in conjunction with either the MHC-GAL4 or Mef2-GAL4 drivers. Two-day-old adult progeny were imaged using a Leica M165FC fluorescent stereo microscope and a Leica EC3 digital camera.

## Protein Quantification

Virgin MHC-, Mef2-, UH3-, and Act88F-GAL4 female flies were crossed with male flies harboring the UAS-Act57BGFP.WT transgene. To quantitate the amount of Act57BGFP.WT transgenic actin driven by each muscle-specific driver, two whole thoraces of, or IFMs from three resulting progeny were dissected and homogenized in Laemmli Sample Buffer (Bio-Rad Laboratories). Each biological sample was then incubated briefly at 100◦C and increasing amounts of protein from each sample were loaded on a 4–15% SDS-PAGE gel (Bio-Rad Laboratories), electrophoresed, and blotted onto a nitrocellulose membrane using the Trans-Blot R TurboTM Transfer system (Bio-Rad Laboratories). Membranes were blocked with gentle shaking in PBS odyssey blocking buffer (LI-COR Biosciences) for one hour, and incubated with primary rabbit anti-actin (Proteintech), goat anti-GFP (R&D), and goat anti-GAPDH (Genscript) antibodies overnight at 4◦C. The membranes were rinsed three–four times, 10–15 min each in 1x TBST (1X TBS with 0.1% tween-20) and then probed with Donkey anti-rabbit and Donkey anti-goat IRDye secondary antibodies (LI-COR Biosciences) for 60–90 min at room temperature. The membranes were rinsed again two–three times, 10–15 min each with 1X TBST followed by a final rinse in 1X TBS. The membranes were subsequently scanned using an Odyssey Infrared Imager (LI-COR Biosciences) (λ = 700 and 800 nm) and analyzed using Odyssey Application Software (v3.030, LI-COR Biosciences). Thoracic and IFM protein quantification was performed on five or eight independent biological samples respectively, with six or four technical replicates each. Mean values (± SEM) of actin and GFP intensities normalized to respective GAPDH intensities were determined. Significance was assessed via One-Way ANOVA with a Bonferroni's multiple comparison test for thoracic, and via the Mann-Whitney test for IFM samples using GraphPad Prism5.

## Flight Testing

Flight tests were performed as described by Drummond et al. (1991). Newly eclosed male and female flies were aged for two days at 25◦C. Each fly was released into the center of a plexiglass chamber with a light source positioned at the top at 23◦C and assigned a flight index of six for upward flight, four for horizontal, two for downward, or zero for no flight. The average flight index from 100–300 flies was calculated for each genotype. Flight assays for Act88F-GAL4-expressing flies were conducted exclusively on females as males consistently displayed severe flight impairment. Values represent mean ± SEM. Significance was assessed using a Kruskal-Wallis One-Way ANOVA.

## Climbing Assay

Climbing tests were conducted on two-day-old flies at room temperature. Small groups of ∼20 flies were placed in covered, cylindrical vials (2.5 cm diameter × 20 cm high), which were aligned with one centimeter markings to measure the height each fly climbed in five seconds. The test was repeated 10 times for each set of flies. The average climbing distance for each fly was recorded for 30–160 flies per genotype. Values represent mean ± SEM. Significance was assessed using a Kruskal-Wallis One-Way ANOVA.

## Imaging of Indirect Flight Muscles

Polarized light microscopy of hemi-thoraces to examine the gross morphology of two-day-old Mef2-GAL4> UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, or UAS-Act57BK326Q/K328Q adult Drosophila IFM was performed as described previously (Nongthomba and Ramachandra, 1999). Briefly, flies were anesthetized, and heads and abdomens removed. Thoraces were fixed overnight in 4% formaldehyde at 4◦C and rinsed in PBS the following day. The specimens were laid supine on a glass slide and snap frozen in liquid nitrogen for 10 s. Frozen thoraces were immediately bisected down the midsagittal plane using a razor blade and IFMs were visualized using a Leica DM5000B microscope at 10X magnification with polarizing filters. Images were taken with a Hamamatsu digital camera.

Fluorescent microscopy was employed to improve pathohistological characterization of IFMs from young (< four hour old) and two-day-old adult Act88F-GAL4> UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, and UAS-Act57BK326Q/K328Q Drosophila. Thoraces were prepared and bisected as described above, followed by staining with 1:100 Alexa-594 Phalloidin in PBST overnight at 4◦C. Samples were rinsed with PBS before imaging with the EVOS R FL Cell Imaging System (Life Technologies) at 4X magnification.

## Results

## Actin Sequence Analysis

The genomes of human, guinea pig, and fly contain six highly conserved actin genes. As found in vertebrates, D. melanogaster expresses specific isoforms of actin in adult skeletal and cardiac muscle. The Actin88F (Act88F) gene encodes all sarcomeric actin of Drosophila IFM (Fyrberg et al., 1983; Hiromi and Hotta, 1985; Nongthomba et al., 2001) while Actin57B (Act57B) is one of two genes encoding sarcomeric actin in the adult fly heart (Cammarato et al., 2011; Shah et al., 2011). The skeletal and cardiac actin isoforms differ in only a few amino acids within and between species (**Figure 2**).

## Generation of Transgenic Drosophila and Confirmation of Muscle-Restricted Transgene Expression

To determine the consequences of masking the inherent charge of actin residues K326 and K328 in vivo, we generated multiple transgenic acetyl-mimetic lines that permitted muscle targeted gene expression. Use of the PhiC31 integrase system ensured all UAS-Act57B transgenes were integrated at an identical, predetermined genomic location (Groth et al., 2004). Thus, our results were directly comparable and any phenotypic differences in control vs. mutant flies could be directly attributed to neutralized lysine charges on the ectopically expressed actin.

To confirm transcription of transgenic actin, flies with constructs consisting of an upstream activating sequence (UAS) followed by a downstream Act57BWT , Act57BK326Q, Act57BK328Q, or Act57BK326Q/K328Q transgene were crossed with flies carrying the GAL4 transactivation gene under the control of the Mef2 promoter (Ranganayakulu et al., 1996). The progeny inherit both genes and express the UAS-Act57B transgenes exclusively in musculature. Total RNA was isolated from the thoraces of young (< two days old) flies of each genotype. First-strand cDNA was synthesized followed by Act57B cDNA amplification. The amplified cDNA contained nucleotide sequences unique to the Act57B alleles, confirming transcription of the WT, K326Q, K328Q, or the K326Q/K328Q Act57B cardiac actin transgene (**Figure 3**). To rule out the possibility of contaminating endogenous Act57B cDNA originating from non-IFM thoracic musculature, amplified cDNA from the dissected IFMs of Mef2- GAL4> UAS-Act57BWT flies was also sequenced, which verified the presence and transcription of Act57BWT in Act88F-exclusive musculature (not shown).

To visualize muscle-restricted expression of UAS-Act57B transgenes, in vivo, flies carrying the UAS-Act57BGFP.WT construct were crossed with flies harboring either the MHCor Mef2-GAL4 muscle-specific drivers. MHC-GAL4-driven transgenic Act57BGFP.WT was readily observed in the thoracic musculature, which is predominantly comprised of 13 pairs of relatively large IFM fibers (**Figure 4**). Mef2-GAL4-driven Act57BGFP.WT, however, was detected far more extensively, in most somatic musculature.

## Relative to MHC-GAL4, Mef2-GAL4 Drives Elevated Expression Levels of Transgenic Actin

The PhiC31 integrase system for transgenic fly production limits transgene expression variability and, when used in conjunction with the GAL4-UAS system, permits the onset and magnitude of expression to be manipulated via distinct drivers (Brand and Perrimon, 1993; Groth et al., 2004; Goentoro et al., 2006). To quantify differences in transgenic protein abundance, dissected thoraces from flies expressing Act57BGFP.WT actin by either MHC- or Mef2-GAL4 drivers, as well as from control "non-driven" flies, were subjected to SDS-PAGE, transferred to nitrocellulose, and probed for actin and for GFP (**Figure 5A**). The resulting signal intensities from GFP and actin were measured and normalized to that from GAPDH (**Figures 5B,C**). As expected very little GFP was distinguished among the various controls. Thoracic muscles from MHC- and Mef2-GAL4> UAS-Act57BGFP.WT flies, however, exhibited detectable amounts of GFP-actin. Mef2-GAL4 induced significantly higher expression levels of the GFP-tagged actin in the thoracic musculature compared to MHC-GAL4. The resulting normalized signal was greater than two-fold higher than that determined for MHC-GAL4> UAS-Act57BGFP.WT flies (1.66 ± 0.41 vs. 0.69 ± 0.07). These findings were corroborated using dissected IFMs, which displayed a normalized Act57BGFP.WT signal that was roughly three-fold higher for Mef2-GAL4> UAS-Act57BGFP.WT compared to MHC-GAL4> UAS-Act57BGFP.WT fibers (6.04 ± 1.85 vs. 1.73 ± 0.37). The thoracic or IFM actin/GAPDH ratio did not differ significantly among the genotypes tested (**Figure 5C**). Estimation of signal intensities from the protein bands, detected exclusively with the anti-actin antibody, suggested that Mef2- GAL4> Act57BGFP.WT comprises ∼10–20% of total thoracic actin (not shown).

## GFP-labeled and Acetyl-Mimetic Cardiac Actin Depress Drosophila Flight Ability and Climbing Performance

Flight tests were performed on two-day-old MHC- and Mef2- GAL4> UAS-Act57BGFP.WT , UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, or UAS-Act57BK326Q/K328Q transgenic Drosophila lines to determine if expression of wildtype or acetylmimetic cardiac actin can support IFM function. The driver lines, and the progeny of each driver line crossed to w 1118 flies, served as additional controls. MHC- and Mef2-GAL4 Drosophila by themselves, and the offspring of the driver lines crossed to w 1118, showed normal flight indices (FI = 5.69–5.83) in accord with published values calculated for wildtype flies (**Figure 6A**) (Drummond et al., 1991; Swank et al., 2003; Suggs et al., 2007;

fragments.

musculature of Mef2-GAL4> UAS-Act57B transgenic flies. The

Cammarato et al., 2008; Wang et al., 2012). "Low dose" ectopic expression of all UAS-Act57B actin constructs by MHC-GAL4 had no effect on flight ability. All lines performed equally well (FI = 5.46–5.78). Thus, MHC-GAL4-driven GFP-tagged, wildtype, or acetyl-mimetic Act57B actin supported flight.

Similar to previous results (Röper et al., 2005), Mef2-GAL4> Act57BGFP.WT expression eliminated flight ability (FI = 0.03 ± 0.01) (**Figure 6A**). Interestingly, relatively "high dose" expression of Act57B cardiac actin that lacked the GFP fusion tag had no observable effect on flight as Mef2-GAL4> UAS-Act57BWT Drosophila demonstrated wildtype-like flight ability (FI = 5.62 ± 0.04). Unlike what was found in combination with MHC-GAL4, expression of UAS-Act57BK326Q by Mef2-GAL4 had a slight, but significant effect on the average flight value (FI = 4.99 ± 0.09) whereas UAS-Act57BK328Q and UAS -Act57BK326Q/K328Q expression abolished IFM function (FI = 0.11 ± 0.02 and 0.08 ± 0.04, respectively). Notably, Mef2-GAL4-driven UAS-Act57BK326Q/K328Q actin expression predominantly resulted in pupal lethality with relatively few adult flies emerging from their puparia. Thus, Mef2-GAL4-driven GFP-tagged or acetyl-mimetic Act57B actin impaired flight and muscle performance.

To assess if expression of acetyl-mimetic cardiac actin affected additional somatic musculature, the climbing ability of w 1118 , Mef2-GAL4> UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, and UAS-Act57BK326Q/K328Q flies was examined (**Figure 6B**). Expression of UAS-Act57BWT actin had no significant effect on climbing distance (8.33 ± 0.54 cm) relative to w 1118 controls (8.85 ± 0.49 cm). All acetyl-mimetic actinexpressing flies, however, exhibited climbing defects compared to w 1118 and to Mef2-GAL4> UAS-Act57BWT Drosophila. Flies expressing UAS-Act57BK326Q had a small but significant reduction in climbing ability (6.14 ± 0.50 cm), while flies expressing UAS-Act57BK328Q or UAS-Act57BK326Q/K328Q had strikingly reduced climbing capabilities (4.22 ± 0.30 cm and 0.93 ± 0.24 cm respectively) compared to controls. Furthermore, flies expressing UAS-Act57BK326Q/K328Q actin performed significantly worse than those expressing UAS-Act57BK328Q actin.

## Mef2-GAL4> UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q Lack of Flight is Associated with Loss of IFM Fibers

In addition to inducing a lack of flight, Mef2-GAL4> Act57BK328Q and Act57BK326Q/K328Q acetyl-mimetic cardiac actin expression also resulted in a "wings up" phenotype. This phenotype is commonly associated with "hypercontracted" and damaged IFM that occurs due to mutations in Drosophila muscle proteins (Fyrberg et al., 1990; Beall and Fyrberg, 1991; Kronert et al., 1995; An and Mogami, 1996; Reedy et al., 2000; Nongthomba et al., 2003). Polarized light microscopy was employed to investigate disturbances in gross IFM morphology (**Figure 6C**). Mef2-GAL4> UAS-Act57BWT and UAS-Act57BK326Q Drosophila displayed similar IFM fiber structure. Both perpendicularly-oriented sets of fibers were continuous and straight, spanning the entire thorax in each line. However, Mef2-GAL4> UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q Drosophila were characterized by the absence of continuous IFM fibers. Only occasionally was light observed emerging from birefringent material closely associated with the thoracic cuticle, which we assumed were IFM fiber remnants. Interestingly, Mef2-GAL4> UAS-Act57BK328Q; Mhc10/+

Drosophila displayed a greater abundance of thoracic material and birefringent musculature consistent with previous studies that demonstrated a reduced number of myosin motors can suppress mutant fiber destruction (Beall and Fyrberg, 1991; Nongthomba et al., 2003). However, the extent of suppression was incomplete as IFMs were still largely absent.

replicates each, and averaged for each genotype. Mef2-GAL4 drove significantly higher amounts of transgenic actin relative to MHC-GAL4

## Relative to UH3-GAL4, Act88F-GAL4 Drives Elevated Expression Levels of IFM-Restricted Transgenic Actin

IFM-specific GAL4 drivers provide an additional resource to further characterize the effects of pseudo-acetylated cardiac actin on flight muscle function and morphology. To quantify differences in transgenic protein abundance exclusively in IFMs, dissected fibers from flies expressing Act57BGFP.WT driven by either UH3- or Act88F-GAL4, as well as fibers from control "nondriven" flies, were subjected to quantitative western blot analysis (**Figure 7A**). The signal intensities from the GFP and actin bands were measured and normalized to that from GAPDH. As found in whole thoraces, GFP signals in IFMs from all control lines were negligible (not shown), while IFMs from UH3- and Act88F-GAL4> UAS-Act57BGFP.WT flies showed detectable amounts of GFP-actin. Act88F-GAL4 induced significantly higher expression levels of GFP-actin in the IFMs compared to UH3-GAL4 (10.31 ± 2.94 vs. 1.57 ± 0.52). No differences in the relative abundance of non GFP-labeled, endogenous IFM actin were identified (not shown).

## IFM-Specific Expression of GFP-labeled or Acetyl-Mimetic Actin Decreases Flight Ability in a Dose-Dependent Manner

Flight tests were performed on two-day-old UH3- and Act88F-GAL4 driver lines and on the progeny of each driver line crossed to w 1118 control, UAS-Act57BGFP.WT , UAS-Act57BWT , UAS-Act57BK326Q, UAS-Act57BK328Q, or UAS-Act57BK326Q/K328Q transgenic Drosophila (**Figure 7B**). The UH3-GAL4 line, and the progeny of UH3-GAL4 crossed to w 1118 displayed unperturbed flight ability (FI = 5.88 ± 0.03 and 5.72 ± 0.05, respectively). "Low dose" expression of all UAS-Act57B actin constructs using the UH3-GAL4 driver had no effect on flight ability. All lines displayed flight indices similar to controls (FI = 5.42 – 5.61). Therefore, UH3-GAL4-driven GFP-tagged, wildtype, or acetyl-mimetic Act57B permitted flight.

Act88F-GAL4 (88F2) Drosophila displayed markedly reduced flight ability (FI = 1.31 ± 0.12) (**Figure 7B**). This was consistent with impaired flight phenotypes observed with publicly available Act88F-GAL4 lines (w ∗ ; P{Act88F-GAL4.1.3}3 and w ∗ ; P{Act88FGAL4.1.3}81B, P{Act88F:GFP}2/ SM6b) (not shown). When crossed to w 1118 control flies however, female progeny harboring a single Act88F-GAL4 (88F2) gene exhibited wildtype-like flight performance (FI = 5.52 ± 0.07). In contrast, no progeny from the two publicly available Act88F-GAL4 lines, when crossed to w 1118 control flies, regained flight ability (not shown). Therefore, we exclusively tested and compared flight performance of the female offspring of Act88F-GAL4 (88F2) Drosophila crossed to each UAS-Act57B actin transgenic line.

"High dose" Act88F-GAL4> UAS-Act57BGFP.WT expression eradicated flight (FI = 0.04 ± 0.02) (**Figure 7B**). Notably, Act88F-GAL4> UAS-Act57BWT flies demonstrated wildtypelike flight ability (FI = 5.25 ± 0.09). As with Mef2- GAL4, expression of UAS-Act57BK326Q via Act88F-GAL4 slightly depressed flight ability, which closely approached statistical significance. However, Act88F-GAL4-mediated expression of UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q caused a complete loss of flight (FI = 0.04 ± 0.01 and 0.03 ± 0.01, respectively).

## The Act88F-GAL4> UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q Flightless Phenotype is Associated with Hypercontracted IFM

Fluorescent microscopy was employed to inspect the IFM histopathology in the acetyl-mimetic relative to control flies

( ∗P < 0.05).

(**Figure 7C**). Two-day-old Act88F-GAL4> UAS-Act57BK326Q Drosophila did not show obvious differences in fiber morphology compared to Act88F-GAL4> UAS-Act57BWT. Similarly-aged Act88F-GAL4> UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q Drosophila, in contrast, showed prominently hypercontracted IFM fibers, characterized by separation and accumulation of fiber material at one or both attachment sites as previously observed in other Drosophila muscle mutants (Fyrberg et al., 1990; Nongthomba et al., 2003). IFMs in very young UAS-Act57BK328Q expressing adults (< four hour old) had a substantially less severe phenotype (**Figure 7C**). Most fibers could be distinctly visualized with minimal thinning and separation. The results indicated that

controls; #P < 0.01 compared to Mef2-GAL4> UAS-Act57BK328Q) (C)

the hypercontracted phenotype associated with Act57BK328Q, and potentially with Act57BK326Q/K328Q acetyl-mimetic actin, was progressive and deteriorates with age and/or use.

## Discussion

relatively little myosin to hypercontract.

Post-translational modifications represent a means to reversibly or irreversibly alter the physical and chemical nature of proteins. PTMs thereby modulate the molecules' properties and dramatically increase the complexity of biological systems. For example, even a single protein can exist as a diverse mixture of many modified forms (Agnetti et al., 2011). Specific

measurements of a number of PTMs have revealed that several residues of a particular protein can be modified. Moreover, different PTMs may compete with each other for access to a single residue on the same protein. Based on an annotated human database, 62% of cardiac proteins have at least one PTM with phosphorylation dominating, whereas 25% have multiple types of modifications (van Eyk, 2011).

The cardiac thin filament is subject to a host of PTMs that markedly influence the properties of the constituent subunits and can directly affect contractile regulation and muscle performance (Metzger and Westfall, 2004; Sumandea et al., 2004; Agnetti et al., 2011; Solaro and Kobayashi, 2011; van Eyk, 2011). However, the modified status of these proteins is infrequently accounted for in in silico and in vitro experiments. As investigation and discovery of myocardial protein PTMs intensify, determining the in vivo consequences of modifying amino acid residues that lie at highly conserved and at potentially critical locations becomes increasingly important. Establishing model systems that limit genetic diversity and benefit from robust and relatively efficient transgenic tools for organism development and high throughput physiological assessment is imperative.

Here we provide novel data pertaining to recently identified PTMs using Drosophila that express pseudo-acetylated K326Q, K328Q, or K326Q/K328Q cardiac actin in a muscle-restricted manner. Transgenic actin expression was confirmed via thoracic and IFM-specific cDNA analysis and by GFP-based reporter imaging. As previously found with similar GFP-tagged constructs (Röper et al., 2005; Perkins and Tanentzapf, 2014), we observed restricted and repetitive incorporation of transgenic GFP-actin along IFM myofibrils, which, as evaluated by phalloidin labeling, were indistinguishable from wild-type myofibrils (not shown). Röper et al. reported that the IFM was the only muscle in which function was affected by overexpression of GFP-labeled actin (Röper et al., 2005). Muscle-restricted expression of muscle or cytoplasmic GFP-actins using the UAS-GAL4 system was shown to yield flightless adults that were otherwise healthy and fertile. Consistent with earlier studies, it was postulated that impaired IFM function, and in some cases the disrupted flight muscle structure, resulted from an imbalance in the relative amounts of actin and myosin (Beall et al., 1989; Bernstein et al., 1993; Röper et al., 2005; Vigoreaux, 2006). Furthermore, Fyrberg et al. (1998) showed that Drosophila exclusively expressing chimeric actin, consisting of part of Act57B fused with the remaining portion of Act88F in their IFM, exhibited decreased flight ability. Thus, in addition to actin and myosin stoichiometric discrepancies that potentially influence performance, these findings suggest functional non-equivalence of actin isoforms and that the IFM is also exquisitely sensitive to actin sequence variation. However, we observed that "low dose" expression of GFP-actin using the MHC- or UH3-GAL4 driver did not impair flight and that expression of non-GFP-tagged wildtype Act57B actin via MHC-, Mef2-, UH3-, or Act88F-GAL4 muscle-specific drivers had no influence on Drosophila flight ability. Therefore, despite differences in 9 amino acid residues between Act88F and Act57B actin isoforms (**Figure 2**), incorporation of non-mutant cardiac actin into IFM myofibrils appeared to support flight at expression levels dictated by each GAL4 driver. These results imply that the GFP moiety of excessively overexpressed GFP-fused actins directly impairs flight. This may be due, in part, to perturbed Tm movement or myosin crossbridge binding in the IFM consistent with the N-terminal fluorescent protein tag located proximal to Tm and myosin binding sites on actin. Importantly, our findings illustrate that GAL4-mediated transgene expression can be employed to investigate the effects of non-GFP tagged cardiac actin modifications on the readily measurable index of flight.

Compared to UAS-Act57BWT cardiac actin expression, expression of mutant acetyl-mimetic cardiac actin had a dose-dependent effect on flight ability. Moreover, robust expression of UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q via Mef2- or Act88F-GAL4 induced a greater reduction in flight performance relative to UAS-Act57BK326Q. Mef2- or Act88F-GAL4-driven Act57BK326Q also had no resolvable effect on gross IFM morphology. It is conceivable that, rather than a direct effect of K→Q substitution on contraction, the milder phenotype associated with Act57BK326Q might be attributable to reduced actin monomer incorporation, owing to a decrease in protein stability or increase in protease accessibility caused by the mutation. Though difficult to rule out confounding effects on protein stability, the preponderance of buttressing evidence suggests that Act57BK326Q exerts direct yet modest effects on contractile function. Namely, in addition to slightly depressing flight ability, Mef2-GAL4> UAS-Act57BK326Q flies exhibited significantly reduced climbing ability. Moreover, Mef2-GAL4> UAS-Act57BK326Q/K328Q flies displayed impaired climbing relative to Mef2-GAL4> UAS-Act57BK328Q flies. The latter results also highlight an influence of pseudo-acetylated actin on non-fibrillar adult somatic muscles.

The absence of flight in Act57BK328Q and Act57BK326Q/K328Qexpressing flies was associated with a "wings up" phenotype and a loss of IFM fibers. Mutations in Drosophila muscle proteins frequently produce a phenotype referred to as hypercontraction (Fyrberg et al., 1990; Beall and Fyrberg, 1991; Kronert et al., 1995; An and Mogami, 1996; Reedy et al., 2000; Nongthomba et al., 2003). This degenerative syndrome is characterized by muscles that begin to develop normally, and then auto-destruct in an apparently myosin-dependent manner. The IFM of the troponin I and T mutants, hdp2 and up101, respectively, initially develop normally and begin to show signs of degeneration 78 h after puparium formation, concomitant with initial muscle twitching in the developing imago (Naimi et al., 2001; Nongthomba et al., 2003). By the second day of adult life very few sarcomeres remain (Beall and Fyrberg, 1991). The onset of hypercontraction suggests that it is a result of muscle activation, and is not due to abnormal development (Nongthomba et al., 2003).

Both hdp2 and up101 thin filaments exhibit aberrantly positioned Tm in the absence of Ca2+, which results in exposed myosin binding sites at rest (Cammarato et al., 2004; Viswanathan et al., 2014). Consequently, IFM hypercontraction is believed to result from excessive actomyosin interaction and unregulated force production. Models of the F-actin-Tm interface reveal K326 and K328 of actin participate in vital intermolecular electrostatic associations with Tm to establish an energetically favorable conformation (Brown and Cohen, 2005; Li et al., 2011; Barua et al., 2012, 2013; Lehman et al., 2013). Here, Tm is located in an azimuthal location that occludes myosin binding sites on F-actin. Moreover, a recent model of the F-actin-Tm-myosin interface reveals K328 on actin can also directly interact with strongly bound myosin heads (Behrmann et al., 2012). Our data suggest that sufficiently high acetylation of these actin residues can weaken actin-Tm interaction and disrupt the ability of Tm to properly block crossbridge formation and force transmission to the thin filament. Therefore, as with the aforementioned troponin mutations, the acetyl-mimetic actin may similarly trigger hypercontraction. Interestingly, the effect of modifying K328 triggered more severe defects relative to K326, which indicates K328 may be most essential for proper relaxation, in vivo. Moreover, since K328Q actin may also impair strong S1 binding, which is predicted to oppose hypercontraction, our data imply the effects of potentially weakening S1-actin association are less harmful than those that influence Tm positioning, as the muscles still hypercontract.

The importance of these actin residues in thin filament regulation, and the effects associated with potential loss of charge, are further underscored by naturally occurring mutations. For example, the K326N nemaline myopathy actin mutation, which differs from the K326Q acetyl-mimetic isoform investigated here by a single methylene bridge, was reported in patients with stiff muscles and spontaneous contractures, suggesting a hypercontractile phenotype (Jain et al., 2012). Computation of the energy landscape for this mutant revealed reduced actin-Tm interaction energy, which would facilitate a shift of Tm away from myosin binding sites, and would explain the increased Ca2+-sensitivity and hypercontractility of affected muscles (Jain et al., 2012; Orzechowski et al., 2014). Considering the severity of the effects accompanied by high expression of the K328Q mutation observed in the current study, similar charge loss at K328 may not be well-tolerated in higher organisms.

Based on F-actin-Tm, F-actin-Tm-myosin models, and our physiological data, we believe that acetylation of K326 and K328 of actin alters electrostatic associations with Tm and/or myosin, destabilizes Tm's inhibitory position, and thereby enhances actomyosin associations and promotes IFM hypercontraction and muscle destruction. However, our approach does not preclude possible alternative contributors to muscle pathology. For example, the amino acid substitutions may compromise the folding efficiency, thermal stability, and/or polymerization properties of actin filaments as recently observed for particular cardiomyopathy-causing lesions (Mundia et al., 2012; Müller et al., 2012). Thus, increased F-actin and thin filament lability may promote myofilament and sarcomeric degeneration. Additionally, though our data suggest that sufficiently high doses of actyl-mimetic actins are required to elicit dysfunction, we cannot rule out a potential contribution from early transgene activation via the Mef2-GAL4 driver, relative to the others, that disrupts muscle development (Markstein et al., 2008). Thus, the most severe IFM phenotype, which was observed in Mef2-GAL4> UAS-Act57BK328Q and UAS-Act57BK326Q/K328Q Drosophila, may be due to both abundant quantities and premature activation times of transgene expression. If K326 and K328 of actin are required for proper thin filament regulation, excessive myosin binding and force production during early muscle development may alter the core building blocks required for proper IFM formation and irreversibly disrupt myofibrillogenesis. This is not unreasonable since during development actomyosin associations appear compulsory for well-ordered and properly functioning IFM (Cripps et al., 1999). However, we detected a greater abundance of birefringent IFM material in Mef2-GAL4> UAS-Act57BK328Q thoraces when myosin was reduced. Moreover, relative to Mef2-GAL4> UAS-Act57BK328Q, delayed expression of UAS-Act57BK328Q actin via Act88F-GAL4 led to a less severe phenotype characterized by post-eclosion progressive separation and accumulation of fiber material to IFM attachment sites. Thus, we interpret these results as consistent with the previously described myosindependent, degenerative hypercontraction syndrome and not with a complete lack of IFM development (Nongthomba et al., 2003).

Here we provide in vivo confirmation of a requirement for positively charged lysine residues at amino acid positions 326 and 328 on actin for proper thin filament function. We demonstrate how PTMs that sufficiently mask these vital charges can have dramatic consequences on muscle performance and structure. While LC-MS/MS analysis of myofilament enriched subfractions of guinea pig hearts revealed lysines 326 and 328 were acetylated, stoichiometry was not assessed (Foster et al., 2013). Our current data suggest Mef2-GAL4-driven transgenic actin comprises approximately 10–20% of total actin (not shown), while MHC-GAL4 drives significantly less. Therefore, minor changes in a potentially small acetylated myofilamentous actin pool may have substantial repercussions on muscle properties. Masking the charges at K326 and K328 apparently increases muscle function by potentially lowering actin-Tm interaction energy, altering Tm positioning, and perpetually promoting myosin crossbridge formation and contraction. Disproportionately excessive amounts of K326 and K328 acetylation and hypercontractile activity in vertebrate hearts may be deleterious as observed here. Nonetheless, small populations of acetylated K326 and K328 could have beneficial effects under normal conditions that act to augment the contractile properties of muscle. In disease however, particularly afflictions characterized by nutrient excess such as diabetes, elevated acetyl-CoA levels may lead to increased acetylation of these critical actin residues and possibly exacerbate pathology.

Investigating the effects of myofilament protein modifications on distinct Drosophila muscles will facilitate our effort to understand the molecular basis of contractile regulation and, importantly, of potentially tempering myopathic responses. Biochemical, biophysical, and structural studies frequently neglect to account for PTMs of myofilamentous proteins, which can greatly modulate contractile behavior. Thus, to truly comprehend muscle performance in health and disease, consideration needs to be given to these dynamic protein modifications. Drosophila facilitates genetic manipulation of thin filament components and evaluation of the consequences of perturbation. Moreover, since abundant quantities of native IFM thin filaments and actin can be isolated for in vitro studies (Bing et al., 1998; Razzaq et al., 1999; Cammarato et al., 2004; Vikhorev et al., 2010), the models permit hierarchical investigation of the effects of such PTMs on contractile machinery from the molecular through the tissue level. Overall, our findings emphasize the utility of Drosophila as a model system that allows for control of genetic modifiers and environmental factors and that enables efficient targeted design and assessment of molecular and tissue-specific responses to protein modifications in the physiological context of muscle.

## Acknowledgments

Fly stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. The authors thank Dr. Sanford I. Bernstein (San Diego State University) for helpful comments and suggestions on the manuscript. ACBB and WS were supported by NIH/NHLBI T-32 HL-07227. Scientist Development Grants from the AHA (12SDG12060056 to DBF and 10SDG4180089 to AC) and NIH/NHLBI R21HL108052 (to DBF) and NIH/NHLBI R56HL124091 (to AC) also supported this study.

## References

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Viswanathan, Blice-Baum, Schmidt, Foster and Cammarato. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

## Skeletal muscle myofilament adaptations to aging, disease, and disuse and their effects on whole muscle performance in older adult humans

## *Mark S. Miller 1, Damien M. Callahan2 and Michael J. Toth2,3\**

*1 Department of Kinesiology, School of Public Health and Health Sciences, University of Massachusetts, Amherst, MA, USA*

*2 Department of Molecular Physiology and Biophysics, College of Medicine, University of Vermont, Burlington, VT, USA*

*3 Department of Medicine, College of Medicine, University of Vermont, Burlington, VT, USA*

#### *Edited by:*

*Julien Ochala, King's College London, UK*

#### *Reviewed by:*

*Ranganath Mamidi, Case Western Reserve University, USA Lars G. Hvid, University of Southern Denmark, Denmark*

#### *\*Correspondence:*

*Michael J. Toth, Health Science Research Facility 126B, University of Vermont, 149 Beaumont Ave., Burlington, VT 05405, USA e-mail<EMAIL_ADDRESS>
Skeletal muscle contractile function declines with aging, disease, and disuse. *In vivo* muscle contractile function depends on a variety of factors, but force, contractile velocity and power generating capacity ultimately derive from the summed contribution of single muscle fibers. The contractile performance of these fibers are, in turn, dependent upon the isoform and function of myofilament proteins they express, with myosin protein expression and its mechanical and kinetic characteristics playing a predominant role. Alterations in myofilament protein biology, therefore, may contribute to the development of functional limitations and disability in these conditions. Recent studies suggest that these conditions are associated with altered single fiber performance due to decreased expression of myofilament proteins and/or changes in myosin-actin cross-bridge interactions. Furthermore, cellular and myofilament-level adaptations are related to diminished whole muscle and whole body performance. Notably, the effect of these various conditions on myofilament and single fiber function tends to be larger in older women compared to older men, which may partially contribute to their higher rates of disability. To maintain functionality and provide the most appropriate and effective countermeasures to aging, disease, and disuse in both sexes, a more thorough understanding is needed of the contribution of myofilament adaptations to functional disability in older men and women and their contribution to tissue level function and mobility impairment.

**Keywords: myosin, actin, cross-bridge kinetics, single fiber, isometric tension, contractile velocity, sex differences, physical activity**

#### **INTRODUCTION**

Aging, disease, and disuse decrease whole skeletal muscle contractile performance, which reduces an individual's ability to accomplish tasks associated with daily living, eventually leading to physical disability (Guralnik et al., 1995; Janssen et al., 2002; Kortebein et al., 2008). Knowledge of the mechanisms underlying the loss of skeletal muscle performance will aid in the development of suitable exercise and pharmacological countermeasures to forestall or counteract these detrimental changes.

Whole muscle contractile performance has been identified as an important determinant of functional limitations in older adults (Reid and Fielding, 2012). As whole muscle performance is dependent upon the functional character of single muscle fibers (Harridge et al., 1996; D'Antona et al., 2006), which are, in turn, largely determined by the type of myofilament proteins they express and their function (Bottinelli, 2001), alterations in myofilament protein biology may contribute to the development of disability in these conditions. Unfortunately, myofilament properties cannot be discerned from measurements performed at the whole muscle level due to methodological limitations (e.g., estimation of muscle size, muscle architecture), subjective factors (e.g., volitional effort) and the interceding effects of other physiological systems that regulate whole muscle function (e.g., neural, excitation-contraction coupling, connective tissue properties). That is, because myofilaments are the end effectors of muscle contraction, the interceding effects of higher order regulatory factors (i.e., at the cellular, tissue or organ/tissue systems level) can mask variation in myofilament function. Thus, to reliably assess the effects of aging, disease, and disuse on skeletal muscle myofilament biology, a reductionist approach is required, where measurements are obtained at the cellular and molecular levels.

This review will focus on human studies that have examined skeletal muscle myofilament structure and function at the cellular and/or molecular levels. As aging, disease, and disuse may alter muscle structure or function by changing muscle quantity [i.e., cross-sectional area (CSA) and/or amount of mass per unit muscle size] or quality (i.e., performance per unit muscle size), we will specifically address these variables in each condition. The review is organized based upon a continuum of whole muscle performance we have observed in our laboratories, which indicates that aging reduces whole muscle performance, both isometric and isokinetic, with progressively larger reductions associated with decreasing activity level, the presence of acute or chronic disease and finally profound, protracted muscle disuse (**Figure 1**). The reader is referred to several recent reviews of other important aspects of the neuromuscular system that undoubtedly conspire with myofilament adaptations to contribute to functional impairments with aging, disease, and disuse (Raj et al., 2010; Rehn et al., 2012; Reid and Fielding, 2012; Russ et al., 2012; Calvani et al., 2013).

#### **MYOSIN-ACTIN CROSS-BRIDGE (XB) INTERACTIONS**

The formation of the myosin-actin XB is the end effector of muscle contraction. This interaction of two myofilament proteins, myosin and actin, dictate single fiber force and contractile velocity, which can be summarized into simple equations by making several assumptions about the myosin-actin interaction: (1) myosin is either strongly bound to actin producing force or detached from actin producing no force and (2) the XB behaves as a Hookian spring, as detailed (Palmer et al., 2007). In brief, the amount of force that can be produced by a half-sarcomere during isometric contraction can be represented as the number of strongly-bound XBs multiplied by the force generated per XB (Funi) (Huxley, 1957; Brenner, 1988). The number of

strongly-bound XBs at any point in time is a function of the total number of functional myosin heads (N) multiplied by the fraction of time a strongly-bound XB is formed (ton) as a function of total myosin cycle time (ton + toff), where ton is the amount of time myosin is strongly bound to actin and toff is the amount of time myosin is detached from actin. Accordingly, the amount of force that can be produced by the half-sarcomere during isometric tension can be written as: N (ton/(ton + toff)) Funi. Notably, ton/(ton + toff) is commonly called the myosin duty ratio and, although difficult to measure, may be altered under various circumstances, with this effect perhaps best exemplified by differences between the myosin isoforms found in human skeletal muscle (Linari et al., 2004). The force generated per XB may also be altered and can be represented as the elastic stiffness of the XB (kstiff) multiplied by the unitary displacement of the myosin power stroke (duni). Contractile velocity is represented most simply as duni divided by ton, in agreement with single fiber findings showing faster velocities with shorter ton (i.e., higher myosin detachment rate) (Piazzesi et al., 2007). Although controversial, recent experimental and modeling work indicates that velocity is also faster with shorter toff (i.e., higher myosin attachment rate) (Hooft et al., 2007; Walcott et al., 2012), potentially via mechanisms that ultimately shorten ton (Walcott et al., 2012). Considered in this context, aging, disease, and/or disuse could alter skeletal muscle force production or velocity by altering the interaction of myosin and actin, which would ultimately affect whole muscle performance and functional capabilities.

What is the relative importance of XB kinetics (e.g., ton and toff) vs. XB mechanical properties (e.g., Funi and duni) in determining single fiber function? Single myosin molecule studies comparing myosin heavy chain (MHC) isoforms between [smooth from turkey vs. skeletal from chicken (Guilford et al., 1997)] and within [α vs. β cardiac in rabbit (Palmiter et al., 1999) and rat (Sugiura et al., 1998)] species have shown no differences in Funi or duni, indicating that isoform differences in force and velocity are due primarily to alterations in XB kinetics. In support of XB kinetics being the primary determinants of fiber function, frog single fiber experiments within a single isoform show that Funi and duni remain consistent over a range of velocities (Piazzesi et al., 2007). In contrast, when looking across fiber types, human single fiber experiments indicate that both XB kinetics and Funi are responsible for the higher forces generated by faster contracting MHC isoforms (Linari et al., 2004). Similarly, single fiber (Brenner et al., 2012) and single molecule (Capitanio et al., 2006) measurements comparing slow isoforms from one species to fast isoforms from another species indicate faster MHC isoforms have higher XB stiffness (kstiff), most likely resulting in a higher Funi. However, these isoform differences may potentially be due to normal sequence variations as isolated myosin from animal species varying in body size contract at distinctly different velocities (Pellegrino et al., 2003). Overall, these results indicate that XB kinetics play an important role in setting both force and velocity in single muscle fibers, while the role of XB mechanical properties is not clearly defined. Thus, the evaluation of XB kinetics and mechanics as mediators of cellular and tissue level contractile function remains an important area of future study, especially in humans.

Alterations in single fiber force and velocity can also occur via changes in muscle quantity and/or structure. Human skeletal muscle fibers contains three different isoforms (I, IIA, or IIX), with individual fibers expressing either one or multiple isoforms leading to six different fiber types (I, I/IIA, IIA, IIA/IIX, IIX, I/IIA/IIX), with pure MHC I and IIA being the two most prevalent. Parenthetically, an additional embryonic isoform of myosin may be expressed in some physiological/pathophysiological conditions (D'Antona et al., 2003, 2006). A simple shift in the type of MHC isoform expressed in the fiber will produce different tissue level velocity and tension (force per CSA) characteristics, as faster contracting MHCs (I *<* IIA *<* IIX) produce more tension (D'Antona et al., 2003, 2006; Pansarasa et al., 2009; Krivickas et al., 2011). Thus, this myofilament variation can alter whole muscle performance (Thorstensson et al., 1977; Ryushi and Fukunaga, 1986; Harridge et al., 1996). In addition to changes in MHC isoform, reducing the amount of myofilament proteins by having smaller fiber CSA, reduced myofibrillar fractional area (e.g., less myofibril area due to an increase in inter-myofibrillar space or other non-contractile elements), or the removal of thick (myosin containing) or thin (actin containing) filaments would decrease single fiber force production. Myofilament ultrastructure plays an important role in setting contractile performance by providing the framework for XB interactions. For instance, at the fiber level, the amount of isometric force produced is equal to the total number of heads interacting in each half-sarcomere (each half-sarcomere must produce identical forces or the sarcomere will change its length). Thus, removal of myosin heads, either from the ends of the thick filament or randomly throughout the thick filament, or a reduction in the number of thin filaments would reduce the number of heads able to interact in a half-sarcomere and result in lower force production. Although fiber CSA is commonly measured in healthy adults as well as during aging, disuse, and disease, changes in myofilament protein content and ultrastructure have not been routinely examined, especially in combination with contractile measurements.

#### **AGING**

Aging reduces whole muscle contractile performance, in part due to a loss of skeletal muscle mass, but also due to reduced muscle quality (Jubrias et al., 1997; Lindle et al., 1997; Lynch et al., 1999; Morse et al., 2005; Delmonico et al., 2009). This age-related diminution in muscle contraction tends to affect dynamic more than isometric function (Lanza et al., 2003; Callahan and Kent-Braun, 2011; Miller et al., 2013), which suggests skeletal muscle properties are altered in a manner that maintains higher forces at slower speeds of movement. Identification of interventions to halt or reverse age-related alterations in skeletal muscle quantity and quality, therefore, can help to maintain physical function and independent living in older adults.

#### **MUSCLE QUALITY**

Multiple human studies have examined whether aging alters contractile performance at the single fiber and myofilament levels with differing results. Findings are generally concordant across fiber types within an individual study, but age-related adaptations in MHC I and IIA fiber contractile performance (isometric tension and velocity) vary widely across studies, with some showing decreases (Larsson et al., 1997; Frontera et al., 2000, 2008; Krivickas et al., 2001; D'Antona et al., 2003; Ochala et al., 2007; Yu et al., 2007) and others showing that function remains unchanged or even increases (Frontera et al., 2003; Trappe et al., 2003; Korhonen et al., 2006; Reid et al., 2012, 2014; Miller et al., 2013). Notably, longitudinal studies show that single fiber tension and velocity either increase or remain constant (Frontera et al., 2008; Reid et al., 2014), suggesting that older adults do not show continued loss of contractile performance and may, in fact, experience improved myofilament function as a compensatory mechanism to offset whole muscle functional decline.

Although single fiber performance results vary widely, myosin protein function appears to decrease from early (e.g., 20–35 year age range) to the older adult (e.g., 60 years and older). In single fibers, and especially in MHC IIA fibers, aging lowers ATPase (Larsson et al., 1997), slows the time response to changes in fiber length (Ochala et al., 2006, 2007), as well as lengthening ton and potentially toff (Miller et al., 2013), indicating slowing of XB kinetics with age. Using isolated myosin, *in vitro* motility studies show that MHC I actin sliding velocity is decreased (Hook et al., 2001; D'Antona et al., 2003) or unchanged (Canepari et al., 2005) with age, while these same studies agree that MHC IIA actin sliding velocity is unchanged (Hook et al., 2001; D'Antona et al., 2003; Canepari et al., 2005). Thus, in contrast to single fibers, results from studies of isolated myosin suggest that MHC I has the larger aging response of slowing XB kinetics (as slower velocity is indicative of a longer ton and/or toff), while MHC IIA is unaffected. These divergent results between these two assays may be related to their experimental conditions as single fiber experiments are performed with their native regulatory protein content and three-dimensional structure using solutions similar to physiologic conditions, while, in order to specifically determine myosin's role in contractile performance, isolated myosin studies remove the myosin from its fiber structure and are typically performed with unregulated actin (Hook et al., 2001; D'Antona et al., 2003; Canepari et al., 2005). Our laboratory (Okada et al., 2008; Miller et al., 2010) and others (Thedinga et al., 1999; D'Antona et al., 2003) have similarly found divergent results when examining myofilament function in isolated myofilaments preparations vs. chemically-skinned single fiber preparations, including when examining genetically-altered animal models (Palmiter et al., 1999; Wang et al., 2013). Based on this evidence, caution must be exercised when interpreting results from these different experimental preparations. Nonetheless, with this caveat in mind, these results collectively suggest that aging generally decreases XB kinetics.

How would slower XB kinetics alter single fiber contractile performance? We would predict that the slowing of XB kinetics, especially longer ton (Miller et al., 2013), should lead to a decrease in contractile velocity. This conclusion is supported by findings of an age-related reduction in contractile velocity that occurs in concert with indications of slower XB kinetics [reduced ATPase activity (Larsson et al., 1997) and longer time response to changes in fiber length (Ochala et al., 2006, 2007)]. At the same time that decreases in XB kinetics reduce velocity, it could have the reciprocal effect of increasing tension, as slower XB kinetics appear to drive an increase in myofilament stiffness, or improved force transmission, that leads to higher isometric tension (Miller et al., 2013). Similarly, slower XB kinetics in older adults occur with an increase in single fiber stiffness, although their overall isometric tension was reduced (Ochala et al., 2006, 2007), which could be explained by reductions in the overall number of XBs secondary to decreased myosin protein expression (D'Antona et al., 2003). Collectively, these studies indicate that the slowing of XB kinetics with age most likely decreases contractile velocity, but may have the paradoxical effect of improving myofilament stiffness and isometric tension.

When findings indicate almost any alteration with age is possible, most noticeable in single fiber contractile performance, this pattern of differences suggests that factors related to the populations studied or the methods employed may explain variation among studies. As the methodology used and experimental conditions are generally standard across studies, variation in the populations studied likely explains the divergent results across studies. One potential reason behind these varying results is differences in physical activity among the populations studied (**Figure 1**), a factor that is difficult to control for and has commonly not been addressed. Physical activity in older adults is well-known to alter single fiber contractile properties (Trappe et al., 2000, 2001; D'Antona et al., 2003, 2007; Frontera et al., 2003; Parente et al., 2008; Harber et al., 2009; Toth et al., 2012), with higher levels of daily activity or specific exercise training regimens tending to reduce or eliminate age-related differences. Such adaptations are not, however, simply linear throughout the activity spectrum, as immobilization in older individuals has been found to increase velocity (D'Antona et al., 2003, 2007), suggesting that complete loss of weight-bearing activity is characterized by very different adaptations, in keeping with findings from animal studies (Reiser et al., 1987) and muscle unloading due to short duration spaceflight in humans (Widrick et al., 1999). Nonetheless, a modulatory role for physical activity with age would suggest that exercise training may mitigate disability, in part, through a reduction of the deleterious age-related changes in myofilaments.

Another potential confounding factor is differences between men and women in their response to aging. Older women have been found to generate higher isometric tensions in MHC I and IIA fibers (Yu et al., 2007; Miller et al., 2013) and have slower contractile velocities in MHC I (Krivickas et al., 2001; Yu et al., 2007) and IIA (Krivickas et al., 2001) fibers. These results are consistent with the finding of slower XB kinetics being more prominent in both of these fiber types in older women (Miller et al., 2013), as we predict slower kinetics would increase tension and decrease velocity. Sex has also been found to have an effect on force production based upon fiber size, as MHC I and IIA fibers with large CSAs produce less force in women compared to men and MHC I fibers with small CSAs produce more force in women (Frontera et al., 2000). However, other studies have found no statistical differences in the single fiber contractile properties of isometric tension and velocity between older men and women (Trappe et al., 2003; Krivickas et al., 2006), with individual laboratories sometimes showing differing results from study to study (Krivickas et al., 2001, 2006). Such variation among studies may reflect differences in other subject characteristics (e.g., physical activity), the number of fibers being examined [i.e., only 7 MHC IIA fibers in older women (Trappe et al., 2003)], or sample sizes as studies not observing sex differences were smaller (*n* = 16–24) than those observing differences (*n* = 24–38). Notably, a large cross-sectional study (*n* = 71) indicates older mobility-limited females produce higher isometric tension in MHC IIA fibers compared to males (Reid et al., 2012), although a three-year longitudinal study (*n* = 16) from the same laboratory found no sex effects on single fiber performance between healthy and mobilitylimited older adults (Reid et al., 2014). Regardless of the reason for variation among studies, these findings raise the possibility that sex differences in myofilament function may contribute to the discrepancies in age-related changes in single fiber performance among studies.

#### **MUSCLE QUANTITY/STRUCTURE**

In general, studies using individual skinned fibers find no change in MHC I and IIA CSA with age (Frontera et al., 2000, 2003, 2008; Ochala et al., 2007; Hvid et al., 2010; Reid et al., 2012, 2014), although some have found decreases in MHC I (D'Antona et al., 2003), IIA (Larsson et al., 1997), or both (Korhonen et al., 2006). CSA also decreases with inactivity in both fiber types (D'Antona et al., 2003). Moreover, CSA has either no (Frontera et al., 2000; Reid et al., 2012, 2014) or varied response to aging in men vs. women, with older men having larger CSAs than young in MHC I fibers (Miller et al., 2013) and older men (Yu et al., 2007) or women (Trappe et al., 2003; Miller et al., 2013) having smaller CSAs than their young counterparts. An advantage of CSA measurements in single skinned fibers is that they are typically performed at the experimental sarcomere length (usually ∼2.50–2.75μm), providing a consistent standard across fiber types and age groups. However, one caveat to the skinned fiber preparation is that it undergoes swelling, which may complicate age comparisons. When CSA measurements are determined from both skinned fibers used for contractile performance and fresh frozen preparations via histochemistry within the same group of volunteers, the two techniques can produce different results, with histochemistry results finding no change in MHC I fibers with age and decreasing in MHC IIA with age (Korhonen et al., 2006; Miller et al., 2013; Callahan et al., 2014a). However, recent studies suggest that age-related differences in skinned human single muscle fibers are comparable to differences observed with fresh frozen preparations (Hvid et al., 2011). Parenthetically, studies in genetic animal models (Blaauw et al., 2009), where muscle fiber protein content is dramatically altered (e.g., 50% hypertrophy in 3 weeks due to activation of Akt signaling) showed disparate CSA results between skinned and fresh frozen preparations, suggesting that variation in the degree of swelling may occur with alterations in fiber protein content, urging caution with this approach. Studies examining whole vastus lateralis muscle indicate that the agerelated reduction in muscle size is due to fibers being lost and a decrease in CSA, especially in MHC II fibers (Lexell et al., 1988; Lexell and Downham, 1991; Lexell, 1995). In contrast, a recent metaanalysis, which did not include skinned fiber results, indicates CSA is unchanged with age, but is larger with higher physical activity levels (Gouzi et al., 2013). Thus, the variation in CSA with age between the various studies may be due to other confounding factors. Overall, these results indicate that CSA with age either remains unchanged or decreases in MHC II fibers, with alterations possibly affected by sex. Despite potential changes in CSA, recent data indicates that the amount of myofilaments as well as their general structure and stoichiometry is unaltered with age or sex (Callahan et al., 2014a). However, studies do indicate age-related changes in post-translational modifications, such as decreased phosphorylation of the fast isoform of the regulatory myosin light chain (Gelfi et al., 2006; Miller et al., 2013), and in gene expression of protein isoforms, such as an increased gene expression of tropomyosin 2 (Roth et al., 2002). These types of alterations have not been commonly examined in human, but may play a role in functional adaptations as they can affect XB performance.

As the MHC isoform distribution and content can alter contractile performance, several studies have examined the agerelated changes in these properties. MHC isoform distribution remained unchanged in longitudinal studies (Frontera et al., 2008) and when physical activity was matched between young and older volunteers (D'Antona et al., 2007; Miller et al., 2013). However, activity level is generally thought to modify MHC isoform distribution, with lower activity causing a shift toward a faster phenotype, or an increase in MHC IIX (D'Antona et al., 2003, 2007), and higher activity causing a shift toward a slower phenotype, or an increase in MHC I (Korhonen et al., 2006; D'Antona et al., 2007). MHC content also remained unchanged in studies matching for physical activity levels (Trappe et al., 2003; Miller et al., 2013) and decreases with reductions in physical activity and immobilization (D'Antona et al., 2003). Altogether, these studies indicate that the amount or types of MHC are not altered by age, but can be changed in older adults based upon their habitual level of exercise.

#### **HEART FAILURE**

Heart failure represents the final common pathway for most chronic cardiovascular disease, and is characterized by an inability of the heart to pump blood to meet the metabolic demands of the body, or that it can only do so at elevated filling pressures. Accordingly, exercise intolerance is the hallmark symptom of the disorder, leading to high rates of disability (Pinsky et al., 1990). Although reduced cardiac contractile performance undoubtedly contributes to reduced functional capacity, changes in skeletal muscle physiology, including muscle atrophy, weakness and reduced oxidative capacity are well-accepted contributors (Zizola and Schulze, 2013). A more complete understanding of the mechanisms causing muscle dysfunction, therefore, is important for identifying appropriate countermeasures to maintain or improve the whole muscle performance and, in turn, reducing disability in these patients.

#### **MUSCLE QUANTITY/STRUCTURE**

Skeletal muscle atrophy in patients with end-stage heart failure has been recognized for decades (Pittman and Cohen, 1964) and studies have identified reductions in single muscle fiber size in patients, especially in MHC IIA fibers (Mancini et al., 1989; Massie et al., 1996; Szentesi et al., 2005), although others have shown that CSA remains unchanged in stable, ambulatory heart failure patients (Sullivan et al., 1990; Schaufelberger et al., 1995, 1997; Miller et al., 2009). Prior results of reduced CSA may be partially explained by muscle disuse-related atrophy, as few studies have controlled for this confounding factor, or because patients were studied shortly following hospitalization. Interestingly, human studies (Toth et al., 2005; Miller et al., 2009), along with animal models (Van Hees et al., 2007), have shown a phenotype of reduced myosin content/functional XB number at the whole tissue and single fiber levels per unit protein or fiber CSA using biochemical (gel electrophoresis) and mechanical (rigor stiffness) measurements, and this loss is observed in patients even when compared to activity-matched controls (Miller et al., 2009). These studies have also suggested that there was no specific reduction in actin protein or thin filament content with heart failure (Miller et al., 2009). At the ultrastructural level, myosin loss could present as a decreased number of thick filaments, shortened thick filaments or a combination of both adaptations. However, no change in either thick-to-thin filament stoichiometry or thick filament length has been found (Miller et al., 2009). Based on this evidence, we hypothesized that myosin was lost at random throughout the thick filament, which would agree with studies that indicate remodeling of the thick filaments takes place by removing and replacing myosin molecules at random throughout the length of the filament (Wenderoth and Eisenberg, 1987; Franchi et al., 1990). Further confirmation of this pattern of thick filament adaption in heart failure will be difficult, as the resolution of current microscopic approaches is insufficient to rigorously test this hypothesis. Nonetheless, the loss of myosin from muscle fibers has clear functional relevance (discussed below).

On a more macroscopic level, heart failure has long been associated with a shift in skeletal muscle fiber type toward a more fast-twitch phenotype (Mancini et al., 1989; Vescovo et al., 1996; Sullivan et al., 1997; Toth et al., 2005). A shift in MHC isoform distribution of this nature could contribute to reduced exercise tolerance given the differing oxidative potential of MHC I and II fibers and this has been demonstrated in patients (Vescovo et al., 1998). However, this pattern of isoform distribution is also emblematic of muscle disuse (Narici and De Boer, 2011) and heart failure patients expend about half as many calories in volitional activity as healthy controls (Toth et al., 1997), suggesting that this adaptation might be due to inactivity accompanying the disease, rather than the disease process itself. This appears to be the case, as we have recently found no alteration in muscle MHC protein isoform expression in patients when compared to activity-matched controls (Miller et al., 2009), and others have shown no differences in MHC isoform expression between patients and controls matched for peak aerobic fitness (Mettauer et al., 2001). There is limited information regarding shifts in isoform expression of other myofilament proteins. One study showed a shift in myosin light chain and troponin T, I, and C toward a slow-twitch isoform distribution in diaphragm muscle in heart failure patients commensurate with a shift in MHC expression toward the MHC I isoform (Tikunov et al., 1996). However, unlike peripheral skeletal muscle, the diaphragm muscle undergoes different functional adaptations (i.e., increased work due to increased pulmonary resistance vs. muscle disuse in peripheral skeletal muscle). In peripheral skeletal muscle, myosin light chain isoform distributions in MHC IIA single fibers were found to show a trend toward greater expression of MLC 1f in heart failure patients (Miller et al., 2010), but this was not associated with differences in single fiber function. Although an indirect measure, an alteration in Ca2+ sensitivity was noted in MHC IIA fibers, which could reflect adaptations in actin-associated regulatory proteins, albeit this was not directly measured.

#### **MUSCLE QUALITY**

Independent of variation in muscle size, numerous studies have suggested that there are reductions in skeletal muscle contractile function at the whole muscle [i.e., tissue level (Harrington et al., 1997; Toth et al., 2010)], suggesting decreased intrinsic function of muscle that may be due to myofilament deficiencies, impaired excitation-contraction coupling (Reiken et al., 2003) or a combination of the two. Studies in humans and animal models over the past 5–10 years have shed light on the possible involvement of myofilaments in these contractile deficits.

The most prominent quantitative/structural change that would be predicted to contribute to strength losses per unit muscle size is a loss of MHC, which should decrease the total number of functional myosin heads, causing a reduction in single fiber isometric tension. Indeed, recent studies have found reduced tension in both in MHC I and IIA fibers from heart failure patients (Szentesi et al., 2005) and in experimental models (Van Hees et al., 2007). However, studies from our laboratory, in which heart failure patients were matched for both age and physical activity level, unlike prior studies (Szentesi et al., 2005), found no diminution in single fiber tension (Miller et al., 2010), arguing that prior results of reduced tension with heart failure were explained more by muscle disuse and/or aging than by heart failure *per se*. The question then becomes: how is tension maintained in the face of a reduction in the overall amount of myosin/functional XBs? We found that XB kinetics were slowed, manifested most prominently as a longer ton, in MHC I and IIA fibers, in patients compared to controls (Miller et al., 2010), a result that has also been observed in animal models of heart failure (Van Hees et al., 2007). This reduction in XB kinetics, if not compensated for by an increase in toff, would lead to an increase in residence time of myosin in the strongly-bound state and, in turn, more myosin heads bound to actin as a function of total myosin cycle time. This is indeed the case, as we found no evidence for a reduction in strongly-bound myosin-actin XBs during maximal Ca2+ activation in heart failure patients using different techniques (Miller et al., 2009, 2010). Thus, adaptations in myosin kinetic properties compensate for the loss of myosin protein content to preserve isometric contractile strength. In fact, building off of our findings from myofilament ultrastructural measures that suggested a random loss of myosin heads along the length of the thick filament detailed above (Miller et al., 2009), we recently reported the results of modeling studies which suggest that this pattern of myosin loss is associated with an increase in myosin attachment time (Tanner et al., 2014). Thus, we hypothesize that a random removal of myosin from the thick filament alters the load imparted to a strongly-bound XB causing a lengthening of ton, leading to a higher duty ratio and higher tension generation.

Although single muscle fiber tension is maintained, reductions in myosin kinetics in heart failure patients may not be without functional consequences. We would predict that reduced XB kinetics, and increased ton specifically, would reduce contractile velocity (Miller and Toth, 2013). Indeed, recent studies from our laboratory show that this is the case in MHC IIA fibers, where lower XB kinetics were associated with decreased single fiber shortening velocity and, in turn, power output (Callahan et al., 2014b). Thus, while we found no evidence for decrements in single fiber tension that would scale to the tissue level to explain reduced isometric force production (Harrington et al., 1997; Toth et al., 2010), reduced myosin-actin XB kinetics may explain reduced muscle power output during dynamic contractile conditions (Toth et al., 2010). This is noteworthy, as reductions in lower-extremity power output are a primary determinant of physical disability (Reid and Fielding, 2012). Thus, alterations in myofilament properties to yield a reduction in the overall capacity for muscle power output during dynamic activities would mean that any activity would be performed at greater percentage of the muscle's physiological capacity and, in turn, may be more fatiguing. In this context, myofilament adaptations may partially contribute to the subjective sensation of exercise intolerance present in heart failure patients, the primary symptom of the disease.

#### **CANCER**

Cancer has well-known effects on skeletal muscle that have been the subject of intense investigation for decades. The most notable effects of cancer are to reduce overall muscle mass secondary to the profound wasting that occurs with the disease [i.e., cancer cachexia (Fearon et al., 2012)], which has, by far, been studied most extensively. However, atrophy is usually confined to certain cancers (e.g., lung, pancreatic, head, and neck) or the later stages of the disease. In contrast, estimates suggest that up to 90% of patients experience subjective functional deficits, manifested most notably as cancer-related fatigue (Cella et al., 2001). Subjective fatigue is a complex psycho-physiological construct, but likely has some roots in reductions in physiological capacity and a recent study suggests that this may relate, in part, to alterations in myofilament biology (Toth et al., 2013).

#### **MUSCLE QUANTITY/STRUCTURE**

Cancer is primarily thought to influence skeletal muscle through its ability to promote muscle atrophy, and has recently been reviewed in detail (Fearon et al., 2012). Despite the well-known atrophic effects of cancer, there are surprisingly few studies that have examined its effects on single muscle fiber size in humans (Weber et al., 2009), although recent work confirms that the atrophy observed at the tissue level extends to the cellular level (Toth et al., 2013). Although profound reductions in the overall amount of contractile tissue occur, recent ultrastructural studies suggest that atrophy is characterized by stoichiometric reductions in the myofilament content relative to fiber CSA (Toth et al., 2013). In contrast to these anatomic measurements, studies that have evaluated myofilament protein content using biochemical approaches in both pre-clinical models (Acharyya et al., 2004) and patients (Eley et al., 2008) have suggested that atrophy in cancer is associated with a selective loss of the contractile protein myosin, with relative preservation of other contractile proteins. If present, one would predict that such an adaptation would diminish muscle contractile force production (Geiger et al., 2000) by decreasing the number of available myosin heads to form strongly-bound XBs. However, more recent studies in the same animal models, as well as patients with cancer, have found no reduction in myosin protein content (Cosper and Leinwand, 2012; Toth et al., 2013). In fact, one study showed that prior findings of a select loss of myosin were likely a methodological artifact (Cosper and Leinwand, 2012). Notably, MHC isoform distribution also remains unchanged with cancer (Toth et al., 2013). Thus, from a quantitative perspective, cancer reduces overall muscle function primarily through its effect to decrease the overall mass of myofilament protein in muscle via simple atrophy, seemingly without selective loss of the major myofilament proteins myosin and actin. This gross loss of contractile tissue manifests functionally as muscle weakness, which undoubtedly contributes to physical disability in cancer patients.

#### **MUSCLE QUALITY**