Mechanical loading stimulates hypertrophy in tissue‐engineered skeletal muscle: Molecular and phenotypic responses

Abstract Mechanical loading of skeletal muscle results in molecular and phenotypic adaptations typified by enhanced muscle size. Studies on humans are limited by the need for repeated sampling, and studies on animals have methodological and ethical limitations. In this investigation, three‐dimensional skeletal muscle was tissue‐engineered utilizing the murine cell line C2C12, which bears resemblance to native tissue and benefits from the advantages of conventional in vitro experiments. The work aimed to determine if mechanical loading induced an anabolic hypertrophic response, akin to that described in vivo after mechanical loading in the form of resistance exercise. Specifically, we temporally investigated candidate gene expression and Akt‐mechanistic target of rapamycin 1 signalling along with myotube growth and tissue function. Mechanical loading (construct length increase of 15%) significantly increased insulin‐like growth factor‐1 and MMP‐2 messenger RNA expression 21 hr after overload, and the levels of the atrophic gene MAFbx were significantly downregulated 45 hr after mechanical overload. In addition, p70S6 kinase and 4EBP‐1 phosphorylation were upregulated immediately after mechanical overload. Maximal contractile force was augmented 45 hr after load with a 265% increase in force, alongside significant hypertrophy of the myotubes within the engineered muscle. Overall, mechanical loading of tissue‐engineered skeletal muscle induced hypertrophy and improved force production.

Therefore, the determination of appropriate models to further our understanding of the plasticity of skeletal muscle to increased and/or decreased loading is critical to understanding muscle-wasting disease.
Tissue engineered three-dimensional (3D) skeletal muscle models have been developed in vitro from primary rodents (Smith, Passey, Greensmith, Mudera, & Lewis, 2012), cell lines (Player et al., 2014), and human muscle precursor cells (Martin et al., 2013;Mudera, Smith, Brady, & Lewis, 2010;Powell, Smiley, Mills, & Vandenburgh, 2002). Engineered muscles are fabricated by seeding cells on or within an extra cellular matrix (ECM) and placing them under uniaxial tension between two fixed points, allowing the differentiation of highly aligned myotubes capable of force generation, similar to that of native muscle tissue (Huang, Dennis, Larkin, & Baar, 2005;Kasper, Turner, Martin, & Sharples, 2018;Khodabukus, Paxton, Donnelly, & Baar, 2007;Martin et al., 2017). Common ECM materials include fibrin (Huang et al., 2005), type I collagen (Cheema et al., 2005;Cheema, Yang, Mudera, Goldspink, & Brown, 2003;Player et al., 2014;Smith et al., 2012) and laminin (Dennis & Kosnik, 2000), either alone or in combination with Matrigel ™ , which promotes cell growth and differentiation (Powell et al., 2002;Shansky, Del Tatto, Chromiak, & Vandenburgh, 1997). Because type I collagen is the most abundant of these matrices in skeletal muscle tissue (Kovanen, 2002), it is an ideal scaffold for engineering in-vivo-like native muscle tissue. Crucially, such systems enable the coupling of mechanistic molecular outputs with morphological and functional measures within a highly controlled environment, without the methodological and ethical constraints of repeated biopsy sampling or animal sacrifice. As such, tissue-engineered 3D skeletal muscle provides an appropriate system to explore skeletal muscle adaptations to mechanical loading.
Loading of skeletal muscle in vitro was traditionally accomplished through mechanical stretch and was first established in monolayer cultures of avian myotubes (Vandenburgh & Kaufman, 1979;Vandenburgh, Hatfaludy, Karlisch, & Shansky, 1989). In this seminal work, cyclic mechanical stretch of collagen embedded monolayer myotubes resulted in an increase in protein synthesis and myotube diameters and changes in the characteristic of skeletal muscle hypertrophy in vivo. Furthermore, it has been established that mechanical loading of monolayer myotube cultures results in the activation of the mechanistic target of rapamycin (mTOR) signalling cascade (Sasai et al., 2010), which has now been well-established for its importance in regulating both protein synthesis and cellular growth Rommel et al., 2001;Wullschleger, Loewith, & Hall, 2006) and is activated after mechanical loading in humans and animals (Bodine, 2006;Bodine et al., 2001;Goodman et al., 2011;Reynolds, Bodine, & Lawrence, 2002).
Mechanical loading of bioengineered muscle has most frequently been used as a method to condition skeletal muscle during myotube development, enabling enhanced orientation, density, and even sarcomeric assembly of myofibers (Boonen et al., 2010;Moon du, Christ, Stitzel, Atala, & Yoo, 2008;Okano & Matsuda, 1997), resulting in a model more greatly analogous to native skeletal muscle. Because it has been established that during myotube development, mechanical loading of tissue engineered 3D skeletal muscle potently induced insulin-like growth factor-1 (IGF-1) messenger RNA (mRNA) expression (Cheema et al., 2005), mechanical loading of such systems may drive a hypertrophic response, given that IGF-1 is implicated in skeletal muscle growth (Velloso, 2008). Whilst data surrounding mechanical loading in postmitotic myotube cultures in 3D are limited, our laboratory has previously demonstrated that both ramp and static loading over a 60 min period in collagen-based tissueengineered 3D skeletal muscle seeded with C2C12 cells resulted in upregulated gene transcripts associated with muscle growth and remodelling such as IGF-1 and matrix metalloprotease 9 (MMP-9) (Player et al., 2014). In addition, mechanical loading of humancraniofacial-muscle-derived myotubes embedded within a sponge of 3D collagen networks induced significant increases in MMP-2 expression (Auluck, Mudera, Hunt, & Lewis, 2005). However, a more detailed characterization of the molecular and phenotypic hypertrophic effects of mechanical loading in 3D engineered muscle has yet to be fully elucidated.
Thus, the aim of the current investigation was to examine the effect of mechanically loading 3D tissue-engineered skeletal muscle on mediating skeletal muscle hypertrophy. Specifically, progressive mechanical overload of engineered muscle was used to determine its temporal effects on myotube hypertrophy, tissue function, mTOR signalling, and well-known hypertrophic and atrophic transcripts (IGF-1, MMP-2, MMP-9, MAFbx, and MuRF-1). We hypothesised that progressive mechanical overload would induce an increase in specific mechano-regulated gene transcripts and posttranslational responses similar to those observed in vivo, ultimately inducing myotube hypertrophy and augmenting maximal force production.

| Cell culture
The immortalized C2C12 murine skeletal muscle myoblast cell line (RRID: CVCL_0188; ECACC, Sigma Aldrich, UK) was used for all experiments herein, and, as such, this study was exempt from receiving full approval from the Loughborough University ethics committee. C2C12 murine myoblast cells were sub-cultured in T80 flasks (Nunc; Thermo Fisher Scientific, UK) and incubated in a humidified 5% CO 2 atmosphere. Cells were cultured in growth media (GM: high glucose Dulbecco's modified Eagle's media [DMEM, Thermo Fisher Scientific, UK]), supplemented with 20% (v/v) fetal bovine serum (FBS, PAN Biotech, Germany) and 1% (v/v) penicillinstreptomycin (P/S; Gibco, UK). GM was changed every 24 hr until 80% confluence was reached, before passage. For all experimentation, cells had undergone fewer than 12 passages.

| Fabrication of tissue engineering skeletal muscle
The printing of the tissue engineering inserts was performed by fused deposition modelling (FDM). Inserts were designed with a twin or single post positioned at the end of an open-ended rectangular chamber, alongside a matching loading bar ( Figure 1). All the relevant standard tessellation language (.stl) files for the designs contained within this manuscript are freely available to download at the following domain: https://figshare.com/projects/3D_Printed_Tissue_ Engineering_Scaffolds/36494. Chamber sizes were scaled to match the volume of collagen required for each construct (Table 1). All 3D F I G U R E 1 Experimental diagrams of: (a) CAD model of the MSB used for mechanical overload, i) tissue engineered skeletal muscle mould, ii) mechanical loading bar, iii) culture plate border containing three loading portals, iv) movement arm containing two rods attached to loading bar, and v) stepper motor. (b) Graphical illustration of progressive mechanical overload regime [Color figure can be viewed at wileyonlinelibrary.com] T A B L E 1 Internal culture dimensions for each collagen gel insert utilised and left to evaporate. All samples were then further sterilised via UV irradiation for 24 hr before use. 3D skeletal muscle was engineered as previously described (Player et al., 2014).
minimum essential medium (MEM; Gibco, UK) was added to 85% type I rat-tail collagen (First Link, UK; in 0.1 M acetic acid, protein concentration 2.035 mg/ml). The solution was neutralised using 5 M and 1 M sodium hydroxide (NaOH) in a drop-wise fashion until a colour change (yellow to cirrus pink) was observed. Subsequently, 4 × 10 6 cells/ml C2C12 murine skeletal myoblast cells (ECAAC, UK) were added in a total volume of 0.1 ml DMEM (Gibco, UK). The collagen/cell solution was pipetted into each 3D printed custom well insert ( Table 1). The engineered muscles were then placed in a 37°C humidified incubator with 5% CO 2 for up to 30 min to set.
Subsequently, each 500 µl construct was manually detached using a scalpel blade from the base of the mould. Three milliliter of GM was added to each construct. The engineered muscles were then incubated in GM for 4 days, with media replenished daily. After 4 days in culture, GM was removed and replaced with differentiation media (DM: high glucose DMEM [Sigma Aldrich, UK]), supplemented with 2% horse serum (Sigma Aldrich, UK), and 1% P/S (Gibco, UK), which was changed daily for a further 10 days.

| Experimental design
After 14 days of culture, engineered muscles were floated in DM and were progressively mechanically loaded using a mechanical stimulation bioreactor (MSB). The MSB is a device programmed to apply progressive mechanical load ( Figure 1) to several engineered muscles, using a floatation bar attached to two pins mounted to a stepper motor (Parker). The progressive load regime ( Figure 1) described a continuous increasing load to achieve 15% stretch over a 1-hr period; thereafter engineered muscle was left under tension (15% stretch) for a further 2 hr, enabling maximal mechanical load upon the myotubes. Engineered muscles were collected at differing timepoints respective to outcome measures. Non-loaded engineered muscle will be referred to as CON.
RNA concentration and quality were assessed by UV spectroscopy at  Table 2. All reactions were performed in triplicate.

| Immunoblotting
Due to the known scalability of the engineered muscles , 500 µl engineered muscles were utilized due to protein quantity constraints within the 50 µl engineered muscles. Engineered muscles were removed 0 hr, 30 min, 3 hr, 6 hr, and 21 hr after T A B L E 2 Primer sequences for the housekeeping gene POLR2B, IGF-1, as well as MMP-2 and MMP-9 which were used as markers of muscle hypertrophy and the E3 ubiquitin ligases MuRF-1 and MAFbx which were used as markers of muscle atrophy

| Assessment of functional muscle
To determine if mechanical loading produced an increase in maximal tetanic skeletal muscle force, 21 and 45 hr after mechanical loading,

| Statistical analysis
All statistical analysis was performed using SPSS software version 23 (RRID: SCR_002865; SPS Inc., Chicago, IL). Normality of distribution and homogeneity of variance in all data sets were determined using a Shapiro-Wilk test and Levine's test, respectively. Data sets were then appropriately analyzed using a one-way ANOVA with an Fishers Least

Significant Difference (LSD) post hoc test. A Kruskal-Willis test was
performed where data were not normally distributed. All data are presented as mean ± standard deviation (SD).

| RESULTS
3.1 | Regulation of hypertrophic and atrophic related genes following mechanical overload 3D engineered skeletal muscle was mechanically loaded for 3 hr as described in Figure 1, and the mRNA levels of candidate hypertrophy-related genes were examined over the subsequent 48 hr

| Mechanical loading increases the phosphorylation of Akt and mTORC1 targets p70S6K and 4EBP-1
Due to its importance in cellular growth, members of the mTORC1 pathway Akt, p70S6K, and 4EBP-1 were investigated after mechanical loading (Figure 3). Phosphorylation of Akt was significantly upregulated 3, 6, and 21 hr after loading when compared to CON (p < .05), with peak phosphorylation at 3 hr. Thereafter, Akt remained elevated 6 and 21 hr after loading. Peak phosphorylation of both p70S6K and 4EBP-1 was evident at 6 hr after mechanical loading and remained elevated at 21 hr relative to CON (p < .05).

| Increased mechanical loading is associated with myotube hypertrophy in tissue engineered skeletal muscle
To discern whether the apparent augmentation of anabolic gene expression and signalling are translated into muscle growth after mechanical loading of engineered muscle, we next investigated myotube size and related morphology (Figure 4)

| Mechanical loading enhances maximal contractile force production
Maximal isometric force was measured in 3D engineered skeletal muscle after mechanical loading ( Figure 5). Compared to nonloaded muscles, relative force production increased in a step-wise fashion 21 (140%) and 45 hr (265%) after the cessation of stimulation compared to CON (p < .01). By contrast, there was no augmentation of muscle function as a result of the additional time in culture in the absence of mechanical loading (p = 1.00).

| DISCUSSION
Mechanical loading in both humans and animals result in skeletal muscle hypertrophy and increased maximal force production, with both components mediated through alterations in cellular signalling and gene expression. To date, research is lacking an appropriate experimental in vitro model to study mechanical loading. In the present study, 3D tissue engineered skeletal muscle was mechanically loaded using a bespoke bioreactor. Mechanical loading temporally activated mTORC1 signalling, gene expression transcripts associated with skeletal muscle hypertrophy, alongside myotube hypertrophy and increases in maximal force production in the days following mechanical loading.
Previously, 3D engineered skeletal muscle has been mechanically loaded during myoblast differentiation (Boonen et al., 2010;Moon du et al., 2008;Okano & Matsuda, 1997), establishing improvements in myotube development and morphology. Our laboratory has previously mechanically loaded highly differentiated 3D engineered muscle and observed a gene expression profile associated with F I G U R E 3 Phosphorylation of p70S6 kinase, 4EBP-1, and Akt at various time points after mechanical loading. Phosphorylated p70S6 kinase, 4EBP-1, and Akt is normalized to the Gel Code blue stain reagent stain. Data are expressed as mean ± SD for n = 7 engineered muscles. Significant values are identified using * where a significance of p ≤ .05 was achieved. SD, standard deviation [Color figure can be viewed at wileyonlinelibrary.com] muscle growth and regeneration; however phenotypic effects were not investigated (Player et al., 2014). In the present study, we measured the mRNA levels of candidate genes based on literature and our previous observations, demonstrating significant increases in IGF-1 mRNA expression 21 hr after the cessation of loading.
Although increases in IGF-1 mRNA were apparent after loading, the magnitude was somewhat lower than that we have observed in our previous experiments (Player et al., 2014), but similar to the effects of muscle loading in humans (Bamman et al., 2001;Deldicque et al., 2005).
We also examined the mRNA levels of matrix metalloprotease 2 and 9 (MMP-2, 9), both of which are pivotal to skeletal muscle development, repair, and adaptive modifications induced by physical exercise (Carmeli, Moas, Lennon, & Powers, 2005;Rullman et al., 2007). In vivo mechanical loading including those mimicking eccentric (Heinemeier et al., 2007), high impact (Carmeli et al., 2005;Rullman et al., 2007) and endurance exercise (Rullman et al., 2009), have produced large increases in the local production of MMP-2 and MMP-9 in skeletal muscle. In the present study, it was observed that MMP-2 mRNA expression was significantly upregulated 21 hr after loading. However, MMP-9 mRNA was left unaltered, in contrast to our previously published work, in which large increases in MMP-9 mRNA expression were observed (Player et al., 2014). Discrepancies in the gene expression profiles were highlighted maybe due to differences in engineered skeletal muscle size, design (i.e. flotation bar vs fixed pins), differentiation status, and experimental protocol.
Nevertheless, the stimulated increase in MMP-2 mRNA expression after loading is indicative of enhanced matrix remodelling, enabling subsequent myotube formation and hypertrophy (Choi et al., 2002).
MuRF-1 and MAFbx are E3 ubiquitin ligases are understood to act through the ubiquitin proteasome pathway (Glass, 2003;Yang, Jemiolo, Trappe, & Proteolytic, 2006), leading to muscle protein  (Ogasawara et al., 2016). We show here that mechanically loaded engineered muscle exhibits step-wise increases in the phosphorylation of both downstream mTOR targets, with peak phosphorylation 21 hr after the loading period. Whilst we cannot account for differences in the temporal activation of mTORC1 signalling in our system compared to in vivo data, a small but significant increase in the hyperphosphorylation status of p70S6k and 4EBP-1 16 hr after resistance exercise has previously been reported in rats (Kubica et al., 2005). To further examine the nature of mTORC1 activation, we also assessed Akt phosphorylation after mechanical loading. Indeed, our observation that Akt phosphorylation was stimulated by loading suggests that the activation of mTORC1 was, at least in part, due to a growth-factormediated pathway. This is in contrast to specific data in humans where Akt phosphorylation has been shown to decrease after resistance exercise (Deldicque et al., 2008), and deletion of the IGF-1 receptor or the pharmacological inhibition of PI3 Kinase (which are upstream positive regulators of Akt) does not impair loadinginduced mTORC1 activation and muscle growth (Hornberger et al., 2004;Spangenburg, Le Roith, Ward, & Bodine, 2008). Nevertheless, in humans, Akt phosphorylation has also been observed to elevate as early as 60 min after resistance exercise (Dreyer et al., 2006), comparable to the current investigation and remains elevated up to 24 hr after resistance exercise (Mayhew, Kim, Cross, Ferrando, & Bamman, 2009). As such, in our system, both loading and growth factors (presumably IGF-1) likely contribute to the activation of mTORC1.
A noteworthy finding from the present study was that after mechanical loading, myotube fusion, and size both increased, a characteristic of myotube hypertrophy. Mechanical stretch of in vitro bioengineered muscle has previously been shown to improve both the orientation and density of rodent myofibers cultured in collagen gels (Okano & Matsuda, 1997). Yet, mechanical loading of postmitotic engineered muscle and its effects on myotube development are yet to be fully elucidated. We show here that myotube hypertrophy in the days after mechanical loading is associated with increased myonuclear number and increased fusion index. As such, this myonuclear addition resembles the satellite-cell-mediated myogenic response observed in native tissue after exercise (Bruusgaard, Johansen, Egner, Rana, & Gundersen, 2010), and likely contributes, alongside IGF-1-Akt-mTORC1 activation, to the enhancement in myotube size. Finally, we were able to establish enhanced functionality of the mechanically loaded engineered muscle F I G U R E 5 Maximal contractile force from 50 µl engineered skeletal muscle cultures at each time point 21 and 45 hr after mechanical loading. All cultures were compared to CON at day 14 within individual experimental repeats to calculate relative force. Data are representative of three experimental repeats and expressed as mean ± SD, for n = 9 engineered muscles. SD, standard deviation [Color figure can be viewed at wileyonlinelibrary.com] AGUILAR-AGON ET AL.

| 23555
through an increase in maximal contractile force production compared to unloaded controls. Recently, the use of electrical stimulation to enable a more mature adult phenotype and increase functionality in engineered muscle has been established (Khodabukus & Baar, 2012;Khodabukus et al., 2019); however research on the use of mechanical loading and engineered muscle functionality and hypertrophy is limited. Although the present model produced force lower than that previously published (Martin et al., 2017;Shimizu, Genma, Gotou, Nagasaka, & Honda, 2017), mechanical overload of the postmitotic engineered skeletal muscle led to significant increases in maximal tetanic force both 21 and 45 hr post, signifying increased functionality of the tissue-engineered skeletal muscles.
Increases in contractile force in systems such as this can be attributed to several factors such as myotube hypertrophy, maturation, and rearrangements within the actin cytoskeleton (Khodabukus & Baar, 2012;Powell et al., 2002). Whilst we cannot discount multiple factors contributing to enhanced contractile force after mechanical loading, we were unable to depict obvious differences in the cytoskeleton staining pattern between stimulated and control engineered muscles, while myotube size was significantly augmented.
In conclusion, we have described the molecular response and morphological and functional adaptation to mechanical loading in mature tissue engineered skeletal muscle. Altogether, the observed increase in IGF-1 mRNA, phosphorylation of Akt, p70S6K, and 4EBP-1, alongside myotube hypertrophy and an increase in functionality of the engineered muscle clearly display an in vitro model of skeletal muscle hypertrophy, similar to that seen in vivo after mechanical loading. As such, this system is amenable to future investigations concerned with enhancing our understanding of skeletal muscle plasticity in health and disease.

FUNDING INFORMATION
The authors would like to acknowledge Loughborough University, the EPSRC EP/L02067X/1, and MRC Centre for Doctoral Training in Regenerative Medicine for funding and support for this study.