Detyrosinated microtubules modulate mechanotransduction in heart and skeletal muscle

Kerr, Jaclyn P.; Robison, Patrick; Shi, Guoli; Bogush, Alexey I.; Kempema, Aaron M.; Hexum, Joseph K.; Becerra, Natalia; Harki, Daniel A.; Martin, Stuart S.; Raiteri, Roberto; Prosser, Benjamin L.; Ward, Christopher W.

doi:10.1038/ncomms9526

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Published: 08 October 2015

Detyrosinated microtubules modulate mechanotransduction in heart and skeletal muscle

Jaclyn P. Kerr¹,
Patrick Robison²,
Guoli Shi¹,
Alexey I. Bogush²,
Aaron M. Kempema³,
Joseph K. Hexum³,
Natalia Becerra⁴,
Daniel A. Harki³,
Stuart S. Martin⁵,
Roberto Raiteri⁴,
Benjamin L. Prosser² &
…
Christopher W. Ward^1,6

Nature Communications volume 6, Article number: 8526 (2015) Cite this article

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Abstract

In striated muscle, X-ROS is the mechanotransduction pathway by which mechanical stress transduced by the microtubule network elicits reactive oxygen species. X-ROS tunes Ca²⁺ signalling in healthy muscle, but in diseases such as Duchenne muscular dystrophy (DMD), microtubule alterations drive elevated X-ROS, disrupting Ca²⁺ homeostasis and impairing function. Here we show that detyrosination, a post-translational modification of α-tubulin, influences X-ROS signalling, contraction speed and cytoskeletal mechanics. In the mdx mouse model of DMD, the pharmacological reduction of detyrosination in vitro ablates aberrant X-ROS and Ca²⁺ signalling, and in vivo it protects against hallmarks of DMD, including workload-induced arrhythmias and contraction-induced injury in skeletal muscle. We conclude that detyrosinated microtubules increase cytoskeletal stiffness and mechanotransduction in striated muscle and that targeting this post-translational modification may have broad therapeutic potential in muscular dystrophies.

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Introduction

The cytoskeleton is critical for the cellular response to the mechanical environment, as it integrates and transduces mechanical energy to mechanosensitive proteins that generate biological signals. Known collectively as mechanotransduction, these pathways regulate diverse cellular physiology¹ and dysregulation of these pathways is linked to disease².

In striated muscle, microtubule (MT)-dependent mechanotransduction of reactive oxygen species (ROS) and calcium (Ca²⁺) signals has been characterized^3,4,5,6,7. Termed ‘X-ROS’ signalling, this mechanotransduction pathway links the mechanical stress of contraction or stretch to NADPH Oxidase 2 (NoX2) generated ROS signals that target Ca²⁺ channels^3,4,6,7,8,9.

The MT contribution to X-ROS has gained considerable attention in the context of Duchenne muscular dystrophy (DMD), in which a dense and disorganized MT network accompanies severe striated muscle dysfunction^5,10. In the mdx mouse model of DMD, we found that increased MT density^4,6 and cytoskeletal stiffness⁴ correlated with excessive X-ROS that induced detrimental Ca²⁺ signals that are believed to underlie cardiac arrhythmia^6,9 and contraction-induced damage in skeletal muscle⁴.

The MT network is a dynamic structure whose density is regulated by enzymes and binding proteins that affect the equilibrium between MT filament growth and disassembly¹¹. In the mdx mouse, we demonstrated that an acute pharmacological strategy to promote MT disassembly (colchicine) reduced X-ROS and detrimental Ca²⁺ signalling in vitro and proffered therapeutic benefit in vivo, implicating increased MT density in the dysregulation of mechanotransduction in DMD. In light of transcriptional evidence from human DMD samples⁴ indicating an upregulation of the X-ROS pathway, as well as a link between MT density and X-ROS in additional dystrophy models^3,7, we initially proposed that the increase in MT network density was a pathological factor in muscular dystrophy.

Assuming force transmission (F) through the cytoskeleton is a product of the stiffness (k) and deformation (x) of the cytoskeleton (F=kx), we reasoned that alterations in stiffness would regulate the magnitude of mechanotransduction. In this regard, we found detyrosinated α-tubulin^12,13 a potentially attractive target for altering cytoskeletal stiffness. Detyrosination is the cleavage of a C-terminal tyrosine residue by an unidentified tubulin carboxypeptidase (TCP)^14,15, a process readily reversed by tubulin tyrosine ligase (TTL)^16,17. This modification stabilizes MT filaments by limiting disassembly^18,19 and can increase the association of the MT network with other cytoskeletal elements^20,21,22, which could increase cytoskeletal stiffness. Intriguingly, detyrosinated α-tubulin is abundant in disease and experimental conditions in which X-ROS is elevated^3,4,6,7, yet its contribution to cytoskeletal stiffness and mechanotransduction remain undefined.

Here we report that MT detyrosination profoundly regulates mechanotransduction by modifying the mechanical properties of the cytoskeleton. By reducing detyrosination, we decreased cytoskeletal stiffness and increased the speed of muscle contraction and relaxation. Furthermore, these cytoskeletal changes dramatically reduced X-ROS signalling and activation of Ca²⁺ channels in both heart and skeletal muscle. These effects occurred without detectable changes in MT network density, indicating a specific role for detyrosination. Consistent with this reduction in X-ROS, we show that in vivo reduction of detyrosination in a murine model of DMD proffers significant protection from contraction-induced injury in skeletal muscle and workload-induced arrhythmia of the heart, both hallmarks of DMD.

We conclude that detyrosinated MT filaments affect the mechanosensitivity of striated muscle by altering the mechanical properties of the cytoskeleton. Given the limitations associated with colchicine treatment²³, the identification of a specific subset of MTs that regulate cytoskeletal stiffness and mechanotransduction is a potentially viable therapeutic option. As this therapeutic strategy has already been established in cancer clinical trials²⁴, targeting detyrosinated α-tubulin may have broad therapeutic potential for the treatment of muscular dystrophies.

Results

Striated muscles are enriched in detyrosinated MT filaments

Immunolabeling α-tubulin reveals a MT network favouring a fenestrated organization in skeletal flexor digitorum brevis (FDB) myofibres (Fig. 1a; control), and a longitudinal axis distribution in wild-type (WT) cardiomyocytes (Fig. 1d; control), consistent with previous reports^10,25,26. Microtubule detyrosination is a post-translational modification (PTM) occurring post-polymerization, and estimates of total detyrosinated α-tubulin in various cell types range from 16-20% in epithelial cells, to approximately 60% in striated muscle, with a limited fraction (<10%) in the free tubulin pool^27,28. Co-labelling cardiomyocytes and skeletal myofibres with an antibody against detyrosinated α-tubulin demonstrates that this PTM is abundant in WT cells and labels a subset of the total polymerized MT network (Fig. 1a,d).

**Figure 1: Detyrosinated α-tubulin is decreased following parthenolide treatment.**

Parthenolide (PTL) is a sesquiterpene lactone that inhibits the activity of the TCP^29,30,31 and decreases the fraction of detyrosinated α-tubulin in vitro. A 2 hr treatment of isolated FDB myofibres or cardiomyocytes (Fig. 1a,d; PTL) with PTL results in an approximate 60% reduction in the amount of detyrosinated microtubules with no appreciable alteration in the density of the total network (Fig. 1b,e). Western blot verified this decrease in detyrosination, and confirmed that PTL did not alter the total α-tubulin content (Fig. 1c,f). As western blot measures both free and polymerized tubulin, and the majority of tubulin in striated muscle is in the free pool^10,32, immunofluorescence is a more accurate indicator of the status of the polymerized network.

We also examined the effects of PTL on other common MT PTMs, namely, acetylation, polyglutamylation and the formation of Δ2 tubulin, all of which are associated with stabilized MTs with each being attributed to functional specificity in MT network regulation^33,34. Although we detected polyglutamylation in FDB fibres and cardiomyocytes, the levels were unaltered by PTL treatment, as shown by western blot and immunofluorescence (Supplementary Figs 1,2). Acetylation was found in FDB fibres and levels were not altered by PTL, whereas little acetylation was detected in cardiomyocytes. We found Δ2-tubulin nearly undetectable in either cell type (Supplementary Figs 1,2). These results are consistent with a previous study of healthy adult myocytes²⁸ and is suggestive of a unique role for detyrosination. Together, these results suggest that PTL specifically reduces the fraction of detyrosinated MT filaments, independent of filament density, tubulin content or other tubulin PTMs.

In contrast to PTL, taxol stabilizes MTs, increasing both network density and the proportion of detyrosinated MT filaments^27,35. Treatment of FDB myofibres and cardiomyocytes in vitro with taxol (Fig. 1a,d; Taxol) significantly increases MT density and the degree of detyrosination by immunofluorescence (Fig. 1b,e). Western blots (Fig. 1c,f) further show that taxol increased the fraction of detyrosinated α-tubulin without altering total α-tubulin content. Taxol treatment also led to a modest increase in acetylation in skeletal myofibers, no detectable acetylation in cardiomyocytes, and little alteration to other PTMs in either cell type (Supplementary Figs 1,2) suggesting detyrosination as a potential contributor to taxol-dependent effects of MT function.

Detyrosinated MTs regulate mechanical properties of muscle

MT detyrosination increases concurrently with the increased MT density that is suggested to suppress contractility in cardiac development^36,37,38 and the failing heart^39,40, and is dramatically increased in diseased skeletal muscle^4,5,7. However, the distinction between MT network density and MT detyrosination per se, and how each contributes to cytoskeletal mechanics, remains unclear. We therefore examined sarcomere dynamics during paced contractions in cardiomyocytes and skeletal myofibres as a gross measure of the contribution of cytoskeletal properties to cellular mechanics. In cardiomyocytes, PTL treatment had no effect on the resting sarcomere length (Fig. 2a; SL) yet significantly increased the degree of sarcomere contraction (Fig. 2a; red) and the maximal velocity of contraction and relaxation (Fig. 2b; red), findings consistent with decreased cytoskeletal resistance. In contrast, taxol decreased the velocity of contraction and relaxation, consistent with increased cytoskeletal resistance (Fig. 2b; blue) as previously described^41,42. In skeletal muscle fibres, neither resting SL nor the change in SL with contraction was significantly different following PTL or taxol treatment (Fig. 3a). The maximal rate of contraction during 1 Hz stimulation was increased with PTL, yet the relaxation rate was not significantly altered (Fig. 3a). However, the time to peak contraction was significantly increased with PTL and decreased with taxol (Fig. 3a), again consistent with the MT network contributing to cytoskeletal viscoelastic resistance. Measurements of calcium transients in PTL-treated cardiac and skeletal muscle cells indicate that changes in global calcium kinetics cannot explain the altered mechanics (Figs 2c and 3c), leaving a decrease in the viscoelastic properties of the cytoskeleton as the most plausible explanation for the increased contractility.

**Figure 2: Contractile mechanics in cardiomyocytes are regulated by detyrosinated microtubules.**

**Figure 3: Mechanical properties of skeletal muscle are regulated by detyrosinated MT filaments.**

We then directly tested the contribution of α-tubulin detyrosination to cytoskeletal stiffness by assaying near-membrane mechanical properties by atomic force microscopy (AFM)⁴. Differentiated skeletal C2C12 myotubes were used due to their robust adhesion to a substrate that permits paired pre- and post-treatment assessments of elastic moduli (Fig. 4d,e). We confirmed that the effects of both PTL and taxol on detyrosination in skeletal myotubes were similar to their respective effects on adult myofibres and myocytes, as shown via western blot (Fig. 4a,b) and immunofluorescence (Fig. 4c). In AFM studies, acute treatment with PTL significantly reduced, while taxol significantly increased, near-membrane transversal stiffness (Fig. 4f). Taken together, the differences in contractile mechanics and near-membrane stiffness support the conclusion that α-tubulin detyrosination can influence the overall mechanical properties of the cytoskeleton. The effects of PTL were independent of significant alterations in MT network density (see Figs 1b,e and 4a,b), suggesting that detyrosination can, by itself, modulate cytoskeletal mechanics.

**Figure 4: Detyrosinated tubulin affects cytoskeletal stiffness**

Detyrosinated MTs regulate X-ROS mechanotransduction

In our initial characterization of X-ROS, a small, brief stretch (∼10% SL, 5–10 s.) resulted in a burst of ROS in cardiomyocytes⁶, though this same protocol failed to evoke an X-ROS signal reliably in WT skeletal muscle fibres⁴. With refined methods and new fiber attachment technology (Fig. 5a,b), we now show that a repetitive sinusoidal length change (5 μm, 0.5 Hz; Fig. 5c) elicits a significant ROS signal in WT FDB myofibres (Fig. 5d,e; grey). The Nox2 inhibitory peptide gp91ds-TAT abolishes the generation of ROS during stretch (Fig. 5d,e; black), confirming that the ROS signal is indeed X-ROS.

**Figure 5: Mechano-activated ROS production and Ca²⁺ influx are regulated by detyrosinated tubulin.**

Having established X-ROS in both skeletal muscle fibres and cardiomyocytes, we sought to determine the role of detyrosination in its generation. We found that skeletal muscle fibres treated with PTL had a significantly reduced X-ROS (Fig. 5d,e; red). To confirm this pharmacological approach, we overexpressed the highly-specific TTL (ref. 43), the enzyme responsible for the reversal of detyrosination by ligation of the C-terminal tyrosine residue on α-tubulin, by electroporation into FDB muscle. In fibres overexpressing TTL, we found a significant reduction in MT detyrosination (Supplementary Fig. 3) and stretch-dependent ROS generation in vitro (Fig. 5d,e; blue), firmly establishing that detyrosinated MTs promote the mechanotransduction of X-ROS.

In adult dystrophin-deficient skeletal muscle, excess X-ROS hyper-sensitized sarcolemmal Ca²⁺ influx channels to drive Ca²⁺ influx during stretch⁴. To test if X-ROS modulates Ca²⁺ influx in WT muscle, we used a modified manganese (Mn²⁺) quench assay and a near-membrane ratiometric calcium indicator (NM-Indo1) to determine sarcolemmal Ca²⁺ permeability during sinusoidal stretch. Following the replacement of extracellular Ca²⁺ with Mn²⁺ (Fig. 5f,g; grey), the stretch-dependent influx of Mn²⁺ through sarcolemmal cation channels quenched the NM-Indo1 signal. The mechanosensitive channel blocker GsMTx4 inhibited this Mn²⁺ quench (Fig. 5f,g blue), confirming the mechanosensitivity of this pathway. PTL treatment also significantly reduced stretch-dependent Mn²⁺ quench (Fig. 5f,g; red), supporting detyrosinated MTs as critical for the mechano-activation of X-ROS and subsequent calcium influx.

In cardiomyocytes, X-ROS is generated with both stretch and electrically evoked contractions^6,9. PTL treatment significantly reduced contraction-induced ROS production in cardiomyocytes, and eliminated ROS production with sinusoidal stretch (1 Hz, 10% of cell length) (Fig. 6a-c). X-ROS was previously demonstrated to increase the sensitivity of ryanodine receptor (RyR) calcium release channels, resulting in an increased frequency of Ca²⁺ ‘sparks’⁶, elementary calcium release events from the sarcoplasmic reticulum (Fig. 6d,e; control). Consistent with the inhibition of X-ROS, we found that PTL treatment dramatically inhibited Ca²⁺ sparks induced by sinusoidal stretch (Fig. 6e).

**Figure 6: Mechanical stress-induced ROS production and Ca²⁺ sparks in heart.**

To control for non-specific effects of PTL treatment on the ROS-dependent regulation of calcium channels independent of MT and mechanotransduction, we applied H₂O₂ to cardiomyocytes and FDB fibres. H₂O₂ elicited a rapid increase in Ca²⁺ spark activity and Ca²⁺influx (Supplementary Fig. 4), independent of PTL. These data indicate that PTL directly interferes with the mechanotransduction through X-ROS, rather than altering Nox2 or ROS-dependent regulation of downstream targets.

Reducing MT detyrosination protects DMD skeletal muscle

Growing evidence implicates a pathological role for altered MTs in cardiomyopathy^{44,45,46,47,48,49} and the muscular dystrophies^4,6,10. Our group and others have demonstrated that acute pharmacologic disruption of the MT network with colchicine is effective at reducing disease-related signalling and functional pathologies in pre-clinical models^4,6,40,50. However, the toxicity profile of this pharmacologic class^23,51,52, underpinned by the interruption of essential MT network functions other than mechanotransduction (e.g., transport, signalling, metabolism), may limit the therapeutic potential of MT disruption.

Our previous work established that X-ROS drives detrimental Ca²⁺ signalling and pathology in heart and skeletal muscle from a mouse model of DMD (mdx mice), and that there is an increase in detyrosinated tubulin in both skeletal⁴ and cardiac⁶ mdx muscle. We confirm here, using our new sinusoidal stretch protocol, that mdx muscle fibres generate more X-ROS during stretch than WT controls (Supplementary Fig. 5). As in WT cells, acute treatment with PTL specifically reduces MT detyrosination and alleviates this excess X-ROS in mdx fibres (Supplementary Fig. 5). Based on this finding and the above in vitro evidence that reducing detyrosination inhibits X-ROS and its downstream consequences (Figs 5 and 6), we tested whether reducing detyrosination in vivo provided a significant benefit to the mdx mouse. We treated adult mdx mice (n=6) with LC-1 (or dimethylamino-parthenolide)⁵³, a prodrug of PTL with enhanced solubility, or vehicle, once a day for three days. On the last day of treatment, mice from each group were evaluated for their resistance to either skeletal or cardiac stress-induced injury.

Resistance to contraction-induced skeletal muscle injury was assessed in vivo by challenging anesthetized mice with 20 eccentric contractions of the hind limb plantar flexors⁴. The isometric torque of the initial contraction revealed no difference between the vehicle and LC-1 treated group, but subsequent contractions revealed a significant protection against contraction-induced force loss in animals treated with LC-1 (Fig. 7a,b). This level of protection was similar to our previous findings with pharmacologic ablation of the MT network or inhibition of Nox activity⁴. Western blot analysis of the contralateral uninjured muscle revealed that LC-1 treatment had no significant effect on Nox2 (gp91^phox), α-tubulin content or levels of acetylated, polyglutamylated, or Δ2-tubulin, while decreasing the amount of detyrosinated tubulin to approximately WT levels (Fig. 7c, Supplementary Fig. 1)⁴.

**Figure 7: Targeting detyrosinated tubulin *in vivo* reduces contraction-induced force decline.**

We previously showed that ablation of the MT network can reduce detrimental mechano-activated [Ca²⁺]_i signalling in mdx fibres⁴, a hallmark of DMD. As a direct test of the effect of LC-1 treatment on contraction-induced Ca²⁺ signalling dysfunction, single mdx FDB myofibres were isolated from the foot of the uninjured leg of each treatment group, loaded with the fluorescent Ca²⁺ dye Fluo-4-acetoxymethyl (AM) and challenged with a 2 min, 5 Hz train of twitch contractions. Before stimulation, both groups exhibited a similar level of cytosolic fluorescence, however, during the 2 min test period, FDBs from LC-1 treated mdx animals maintained significantly lower cytosolic [Ca²⁺] compared with vehicle-treated mdx fibres (Fig. 7f,g). At the end of the pulse train, the return to resting [Ca²⁺] was significantly faster in LC-1 treated fibres, as [Ca²⁺]_i remained elevated above pre-stimulation values for upwards of 100 s following stimulation in untreated mdx fibres (Fig. 7h). Taken together, these results suggest that reducing detyrosination protects against contraction-induced Ca²⁺ signalling dysfunction and injury in mdx muscle.

Reducing MT detyrosination protects DMD heart

We and others have previously shown that mechanical stress drives detrimental ROS and Ca²⁺ signalling in the mdx heart^6,54 that can trigger arrhythmias^55,56. We next assayed the contribution of the detyrosinated MT network to this pathology in the dystrophic heart. Mdx mice treated with either LC-1 or vehicle for three days were anesthetized, instrumented to record electrocardiogram (ECG) and challenged with the β-adrenergic agonist isoproterenol (0.5 mg kg⁻¹) to increase cardiac work. Immediately following isoproterenol challenge, the resting heart rate increased two- to three-fold in both treatment groups. In vehicle-treated mdx mice (Fig. 8a), arrhythmias became apparent within 10 min and progressed in frequency until the mice succumbed to arrhythmia-related cardiac failure and death (mortality in 12 of 13 animals tested). In contrast, no mice in the LC-1 treatment group (n=6) succumbed to arrhythmia-related death (Fig. 8a). Western blot analysis of the vehicle versus LC-1 treated hearts revealed that LC-1 treatment had no significant effect on α-tubulin, acetylated tubulin, or Nox2 (gp91^phox) content, but significantly decreased the amount of detyrosinated α-tubulin (Fig. 8c, Supplementary Fig. 1e). Consistent with LC-1 acting through X-ROS to exert its effect, protection from arrhythmia-associated death was also conferred by the in vivo inhibition of Nox2 with gp91ds-TAT (n=3/3 survival) or by pharmacologic disruption of the MT network with colchicine (n=5/5 survival).

**Figure 8: *In vivo* targeting of detyrosination in *mdx* heart rescues isoproterenol induced arrhythmia and lethality.**

To directly test the role of MT detyrosination in generating arrhythmogenic calcium signals, we used an established high-frequency stimulation protocol to evaluate Ca²⁺-dependent arrhythmogenic potential in single mdx cardiomyocytes loaded with the Ca²⁺ indicator Fluo-3 (ref. 56). Following a 30 s burst of 3Hz electrical pacing, untreated mdx cardiomyocytes exhibited spontaneous, propagating Ca²⁺ waves linked to arrhythmia formation, as well as a significant increase in resting Fluo-3 fluorescence (Fig. 9a–c). These findings are consistent with RyR-mediated calcium ‘leak’ that underscores arrhythmogenesis in mdx myocytes^6,55,56. PTL treatment significantly reduced the frequency of Ca²⁺ waves and prevented the elevation in resting calcium concentration following high-frequency stimulation (Fig. 9a–c), suggesting that targeting MT detyrosination reduces detrimental calcium signalling and stress-dependent arrhythmogenesis in the DMD heart.

**Figure 9: Parthenolide treatment ablates stress-induced Ca²⁺ waves and leak in *mdx* cardiomyocytes.**

Discussion

We report the novel discovery that detyrosination is a potent regulator of MT-dependent mechanotransduction in striated muscle. Using both pharmacologic and genetic strategies, we show that MT detyrosination correlates with the magnitude of X-ROS signalling in heart and skeletal muscle. As the level of detyrosination affects both the velocity of contraction and cytoskeletal stiffness, we conclude that detyrosination regulates mechanotransduction by altering the mechanical properties of the cytoskeleton.

Iribe et al⁵⁷ first identified the role of MTs in the mechanotransduction of cardiac Ca²⁺ signalling. Subsequent studies identified X-ROS as the proximate activator of these Ca²⁺ signals in both cardiac⁶ and skeletal muscle⁴ by showing the blockage of X-ROS with pharmacologic ablation of the MT network. Here we report that a selective reduction in detyrosinated MTs yields a similar effect on mechanotransduction, while preserving the overall MT network. This was at first a surprising finding, particularly as there is no evidence that detyrosination contributes to the intrinsic stiffness of the individual MT filament per se, and historically, some have viewed this PTM (along with acetylation, polyglutamylation and Δ2-tubulin) as simply a marker of stable microtubules.

However, this view is being challenged, and our study adds to an emerging body of work demonstrating specific roles for detyrosination in the regulation of MT function¹⁴, including the organization of chromosomes in mitosis³¹ and the regulation of Na⁺/K⁺ ATPase activity⁵⁸. We found detyrosination uniquely abundant in healthy muscle compared with other common PTMs (Fig. 1, Supplementary Figs 1,2)²⁸, and PTL and LC-1 selectively reduced this PTM without detectably altering acetylation, polyglutamylation or Δ2-tubulin, allowing the attribution of functional consequences to detyrosination. On the other hand, taxol has complex effects³⁵ as it increases the amount of polymerized MTs, and several tubulin modifying enzymes favour the polymerized form¹⁴. Here taxol increased MT network density, dramatically increased detyrosination, and modestly affected other PTMs, and thus results should be interpreted with this in mind. Separating the contribution of these individual PTMs to specific biological functions will be enabled by the continued identification of tubulin modifying enzymes and molecular tools to manipulate them, including the elusive TCP¹⁴.

How can reducing the fraction of detyrosinated MTs have such significant effects on mechanotransduction? While detyrosination may not alter intrinsic MT stiffness in vitro, it does increase MT stability by inhibiting disassembly via motor-proteins¹⁸. In addition, while MTs can resist compressive loads, the mechanical properties of the MTs alone are insufficient⁵⁹, and likely require cross-linking interactions with other proteins and cytoskeletal elements to confer the requisite mechanical stiffness needed to resist cytoskeletal compression⁶⁰. Recent studies linking functional deficits in heart failure and muscular dystrophy to the overexpression of Microtubule-Actin Cross-Linking Factor 1 (ref. 61) or the cytoskeletal cross-linker plectin⁶² offer support for this idea in striated muscle. Further evidence comes from non-muscle cells, where increased cytoskeletal stiffness is driven by interactions between detyrosinated MTs and intermediate filaments (IFs)^21,22,29. In this context, it is notable that the expression of desmin, the major IF in striated muscle, is elevated in the mdx mouse⁶³, and its deletion provides significant functional benefit⁶³. While the authors did not attribute a rescue of dysregulated mechanotransduction to their results, our current data support this possibility, as increased MT:IF cross-linking offers a parsimonious explanation for the detyrosination-dependent elevations in X-ROS signalling and cytoskeletal stiffness. Finally, detyrosination also regulates the interaction of MTs with CAP-Gly proteins, which may have important consequences for both cytoskeletal organization and cell signalling⁶⁴.

DMD provides an ideal model to test the contribution of detyrosinated MTs in mechanotransduction-dependent dysfunction, as an increase in MT network density and detyrosination correlates with pathology in DMD^4,5,6,10. While previous studies have demonstrated that acute pharmacologic disruption of MT structure can be beneficial in rodent models of DMD and pathologic cardiac hypertrophy^4,32, this strategy is limited by the potential for depressing other central MT functions^23,51,52. Targeting detyrosination is an attractive strategy as it exerts significant effects while sparing the MT network. Here we report that acute in vivo pharmacologic reduction of detyrosination protects the DMD heart from workload-induced cardiac death and skeletal muscle from contraction-induced injury. Our in vitro studies confirm that reducing detyrosination inhibited the detrimental Ca²⁺ signals that underlie these DMD pathologies.

Although we limited our pre-clinical effort in this study to DMD, elevated MT density and detyrosination have been identified in multiple models of muscular dystrophy^4,5,7,8 and pathologic cardiac hypertrophy^6,36, suggesting that these MT alterations are conserved maladaptive responses in disease. Despite the growing evidence for a MT-targeted strategy to slow or halt disease progression in DMD and the failing heart, the move to clinical translation has been slowed by the unavailability of promising therapeutic targets and effective pharmacologic agents. PTL is a sesquiterpene lactone that has demonstrated promising utility as an anti-proliferative agent for treating human cancers²⁴. Given that PTL (as feverfew extract) and LC-1 have advanced to Phase I human clinical trials⁶⁵, compounds such as PTL represent novel lead candidates for MT-centric therapy in striated muscle disease. Beneficially, significant clinical data generated by cancer therapeutics⁶⁵ will be available to guide the translation of SL therapy to striated muscle diseases.

In summary, we present evidence that MT detyrosination regulates mechanotransduction through ROS and Ca²⁺ signals. In diseased muscle sensitive to mechanical stress, inhibiting detyrosination blunts the stress-induced ROS and Ca²⁺ signalling dysfunction that drives pathogenic progression. We conclude that MT detyrosination regulates mechanotransduction in striated muscle and that targeting this PTM may have broad therapeutic potential in the muscular dystrophies.

Methods

All reagents and drugs were purchased from Sigma-Aldrich unless otherwise noted. Gp91ds-TAT peptide was purchased from Anaspec. GsMTx4 was kindly provided by F. Sachs.

Murine models

All animal care and experimental procedures were approved and performed in accordance with the ethical standards set by the Institutional Animal Care and Use Committees of both the University of Maryland and the University of Pennsylvania and by the National Institutes of Health (NIH). Mice (stock numbers 001801 and 000664) were acquired from Jackson Laboratories. Rats (Hsd:Sprague Dawley SD) were purchased from Harlan Labs. Animal cohorts within each genotype were randomly assigned to experimental groups.

Preparation of flexor digitorum brevis (FDB) muscle fibres

FDB muscles from male control rat (WT), as well as male control mice (C57/Bl6; WT) and dystrophin-deficient (C57/10ScSn-Dmd^mdx/J; mdx) mice at 9–11 months of age were isolated as follows: after euthanasia, FDB muscles were harvested bilaterally and placed in 4 mg ml⁻¹ collagenase A. Following incubation at 37 °C for 2 h, the muscles were gently triturated to release individual fibres. Fibres were imaged or fixed within 24 h of isolation.

Cardiomyocyte isolation

Adult C57/Bl6 or mdx mice and Sprague–Dawley rats were terminally anesthetized by injection of pentobarbital, followed by excision of the heart and enzymatic isolation of ventricular myocytes⁶⁶. Hearts were suspended from a Langendorff perfusion column via cannulation of the aorta and perfused with cell isolation buffer containing in mM: NaCl 130, HEPES 25, Glucose 22, KCl 5.4, lactic acid 1, MgCl₂ 0.5, NaH₂PO₄ 0.33, CaCl₂ 0.3, to which insulin (100 micro-units per millimetre), collagenase II (0.66 mg ml⁻¹ Worthington LS004176), trypsin (0.033 mg ml⁻¹ Sigma T0303) and proteinase (0.033 mg ml⁻¹ Sigma P8038) were added. Tissue was then minced and subjected to additional digestion with 0.7 mM CaCl₂ and bovine serum albumin (2 mg ml⁻¹) at 37 °C until myocytes were easily liberated by trituration with a Pasteur pipette. Myocytes were then removed from enzyme and then slowly reintroduced to physiological CaCl₂ in cell isolation buffer without enzymes. Cardiomyocytes were stored in a normal Tyrode solution containing (in mM): NaCl 140, KCl 5, CaCl₂ 1.8, MgCl₂ 0.5, HEPES 5, Glucose 5 and NaH₂PO₄ 0.33. Experiments were performed at room temperature, 22 °C. Due to the relatively greater ease and reproducibility of highest quality cell preparations, rat myocytes were used for all experiments except when mdx myocytes and appropriate murine controls were needed.

Immunofluorescence

Isolated FDB fibres were adhered to glass coverslips coated with Geltrex ECM (Life Technologies). Fibres were fixed at room temperature for 20 min using 4% paraformaldehyde in PBS, followed by permeabilization at room temperature for 15 min with 0.1% Triton X-100 in PBS (ref. 67). Non-specific staining was limited by adding SuperBlock PBS (Life Technologies) for 1 h before the addition of primary antibodies. Primary antibodies (α-tubulin clone DM1A, Sigma; rabbit detyrosinated tubulin, AbCam #48389; acetylated tubulin clone 6-11-B, Sigma; rabbit Δ2tubulin, Novus #NB100-57397; mouse polyglutamylation clone GT335, Adipogen) were diluted in SuperBlock PBS and added to the coverslips for an overnight incubation at 4 °C. Secondary antibodies (Goat anti-mouse Alexa 488, 647 and goat anti-rabbit Alexa 488, 568; Life Technologies; 1:100 dilution) were diluted in SuperBlock PBS and incubated at room temperature for 6 h. Coverslips were mounted on glass slides using ProLong Diamond (Life Technologies) and imaged using a Zeiss 510Live laser confocal microscope (× 63 1.4 numerical aperature plan-apo oil objective).

Isolated cardiac myocytes were fixed and permeablized in solution using 4% paraformaldehyde and 0.1% Triton X-100, both in PBS. Myocytes were then transferred to SuperBlock PBS and allowed to settle on coated coverslips for 1 h. The coverslips were then treated with antibodies, mounted and imaged as above.

Quantitation of α- and detyrosinated tubulin was performed with Volocity (Perkin Elmer). Total pixel density above threshold within a region of interest was calculated for each channel. The average density for each treatment condition and channel were calculated and normalized to the control untreated condition.

Skeletal fibre mechanics and [Ca²⁺]_i transients

Isolated FDB fibres treated (2 h) with PTL (10 μM), taxol (10 μM), at room temperature, suspended in Hepes-buffered Ringer solution containing (in mM): 140 NaCl, 4 KCl, 1 MgSO₄, 5 NaHCO₃, 10 glucose and 10 HEPES (pH 7.3). Following treatment, fibres were placed in a custom rotating glass-bottomed perfusion chamber (Four-Hour Day Foundation) mounted over an inverted microscope (Olympus IX-70, × 40 objective). Contractions were elicited under no-load conditions with field stimulation (0.2-ms square pulse, 1 Hz elicited), while SL was monitored and recorded using a high-speed video SL system (HSVL 901B, Aurora Scientific). Across five successive contractions, velocity of shortening and relaxation as well as the time to peak shortening were calculated. Similarly, PTL or untreated FDBs equilibrated with the Ca²⁺ indicator Indo-1PE (5 μM, 0.1% Pluronic F127, 30 min) were electrically stimulated (1 Hz) and the peak, V_min and V_max of the Indo-1 ratio (405 nm/475 nm) were analysed (950 A, Aurora Scientific).

Myocyte contraction and [Ca²⁺]_i transients

Experiments were performed in custom fabricated cell chambers (Four-hour Day Foundation, Towson, MD) mounted on a ZeissLSM 780 inverted confocal microscope with a × 40 Oil 1.4 NA objective. For measurements of contractility, SL was monitored with a high-speed video camera and Fourier transform analysis. The [Ca²⁺]_i transient was evaluated with Fluo-3 by 15 min incubation with 2 μM Fluo-3-AM ester and 0.01% Pluronic F127. Cells were allowed an additional 10 min for de-esterification and then scanned using a 488 nm argon ion laser in confocal line-scan mode at 1.92 ms per line. Cells were paced to steady state using 1 Hz field stimulation through platinum electrodes. After 20 s of 1 Hz stimulation, all cells showed steady state transients and contractions. At this point, five Ca²⁺ transients and five contractions were assayed. This protocol was repeated 2 × in each myocyte, and data was pooled for analysis.

Myofibre attachment and stretch

All experiments were performed in a custom rotating glass-bottomed chamber (Four-Hour Day Foundation) equipped with bath perfusion and mounted on an inverted microscope (Olympus IX-70, 40 objective) fitted with a DG4 (Sutter Instruments), an IonOptix Myostretcher (IonOptix) system with a 403 A-CF force transducer (Aurora Scientific), and PMT Sub-System (IonOptix). Isolated FDB fibres were suspended in Hepes-buffered ringer as above, supplemented with PTL (10 μM) or left untreated (2 h). Individual fibres were attached between microfabricated glass fibre holders (IonOptix), coated with MyoTak biological adhesive (IonOptix), that were mounted to a force transducer (403A-CF, Aurora Scientific) and a Piezo actuator (PI). The force and positional outputs were recorded at 1kHz on a 600A data acquisition system from Aurora Scientific.

Stretch-induced ROS production was assayed in H₂DFFDA (5-(and-6)-carboxy-2′,7′-dihydrofluorescein diacetate; 5 μM, 30 min; Life Technologies) loaded FDB fibres with a sinusoidal waveform (8% of resting SL) imposed for 20 s. Data were collected offline (500 Hz; 950A, Aurora Scientific) and the rate of ROS generation analysed using the Origin Pro 9.1 (Origin Lab). The Nox2-specific inhibitor gp91ds-TAT (2 μM) was used to verify the source of ROS.

Stretch-induced Ca²⁺ influx was assayed with a modified manganese quench (MnQ) assay. Briefly, cells were pre-loaded with Indo-1 Near-Membrane (NearMem, formerly FIP-18, 5 μM, Teflabs) and transferred and attached to the stretch apparatus described above. Perfusion with normal Ringer was switched to a Ringer where Ca²⁺ was substituted with manganese. Fibres were excited (340 nm), stretched with a sinusoidal waveform as above, and sarcolemmal permeability to Ca²⁺ estimated by the manganese influx and quench of the total emission fluorescence (F_TOT=405+475 nm). The rate of fluorescence decrease was calculated and analysed as above. GsMTx4, a specific inhibitor of stretch-activated channels, was used to confirm the source of influx. For certain experiments, fibres were perfused with 200 μm H₂O₂ to elicit sarcolemmal calcium influx, independent of sinusoidal stretch (Supplementary Fig. 4).

Cardiomyocyte attachment and stretch

Glass microrods were coated with a biological adhesive, MyoTak (IonOptix, Milton, MA). One glass microrod was connected to a force transducer (403A-CF, Aurora Scientific), and the other to a length controller (World Precision Instruments). Myocytes were attached at both ends by gently pressing down with the MyoTak-coated microrod and then lifting the cell from the chamber bottom. Axial stretch was applied by movement of the length controller in response to variable voltage output. SL was monitored with a high-speed video camera (Aurora Scientific). Average SL before stretch was 1.8 μm. Myocytes were subjected to three stretch-release trials, with 30–60 s rest allowed between trials. As each stretch produced a similar change in SL, DFF fluorescence, and Ca²⁺ spark rate, trials were pooled for analysis. WT cells did not commonly display Ca²⁺ waves, and cells with Ca²⁺ waves were discarded from analysis.

Ca²⁺ spark measurements

Cells were loaded with Fluo-4 by 10 min incubation with 3 μM Fluo-4-AM ester and 0.01% Pluronic F127 and allowed an additional 10 min for de-esterification. Cells were scanned using a 488 nm argon ion laser in confocal line-scan mode at 1.92 ms per line. Automated analysis of line-scan images for Ca²⁺ spark location and properties was performed using SparkMaster (ImageJ⁶⁸). For H₂O₂ studies, spark measurements were taken in the same cell before and after acute (<1 min) H₂O₂ (200 μM) exposure. Some cells were pretreated with PTL (2 h, 10 μM).

DFF measurements in cardiomyocytes

Cells were loaded with the cell permeant ROS indicator DFF (a more photostable analogue to the commonly used DCF) by a 20 min exposure to a Tyrode’s solution containing 2.5 μM H₂DFFDA. The cells were imaged using confocal microscopy at 1 frame per second. Cells were imaged at very low laser intensity and with a sampling rest interval of 2 s to avoid artifactual signal amplification on continuous light exposure. The slope of the DFF signal is indicative of ROS production; therefore the slope before stretch has been subtracted from the entire trace so that the initial slope is zero and changes in slope can be easily visualized^4,6. The slope is expressed normalized to the slope before stretch in that same cell as this comparison minimizes differences in DFF loading and resting signal between cells. As the slope was linear for the duration of cyclic stretch, the slope was calculated from the entire stretch duration for all cyclic stretch experiments.

Atomic force microscopy

The transversal stiffness of differentiated C2C12 myotubes was quantitatively measured by means of nanoindentation measurements based on atomic force microscopy (AFM). Murine C2C12 myoblast cell line provides a well-established and reproducible model system for skeletal myogenesis⁶⁹. When 80% cell confluence was reached, the medium containing 10% foetal calf serum was substituted with 2% foetal calf serum to induce myogenic differentiation, a process lasting a few days which can be recognized by a progressive elongation in cell shape and an assembling of structures involved in contraction. Differentiated C2C12 cells (American Type Culture Collection, ATCC) cultured on coverglass were treated for 2 h with PTL, Taxol or dimethylsulphoxide and then fixed and immunostained for MT as described above.

A commercial AFM (model 5500, Keysight Technologies) coupled with an inverted optical microscope was employed to record force-versus-distance curves, taken at a constant speed of 3 μm s⁻¹ on differentiated C2C12 myotubes. At least three 500 × 500 nm² areas, selected over the cell body to avoid nuclei regions, were sampled with a regular grid of 16 × 16 force curves each, for a total number of force curves ≥768 for each considered cell. The same commercial silicon microcantilever with a conical tip (model CSG11, NT-MDT, nominal spring constant k=0.03 N m⁻¹) was used for all measurements. The region after contact of each force curve was transformed into a load-versus-indentation curve and the corresponding contact stiffness could be calculated at a certain maximum indentation. To maximize the sensitivity of the measurement towards the microtubule network structure and organization the reported values were calculated at a maximum indentation of 100 nm. Acute treatments were performed by incubating the already differentiated C2C12 culture for 2 in the medium containing either parthenolide or taxol at a final concentration of 10 μM. Before starting the AFM measurements the solution was replaced with fresh medium. Following AFM indentation measurement, cultures were labelled by using a primary antibody against α-actinin (clone EA-53, Sigma; goat anti-mouse Alexa 549 secondary) and fluorescent phalloidin (Alexa 488) to confirm the periodic sarcomeric organization of striated muscle fibres.

Formulation of LC-1

The parthenolide prodrug LC-1 (ref. 53) was formulated for oral dosing in 2-hydroxypropyl-β-cyclodextrin (HP-β-CD)⁷⁰. In brief, 160 mg of LC-1 were dissolved in 3.75 ml of an aqueous 0.3 M HP-β-CD solution and mixed well. The LC-1 and vehicle only solutions were stored at 4 °C before administration.

In vivo injury

Mdx mice (9–11 months) were treated daily via gavage with a bioavailable parthenolide analogue (LC-1) or vehicle (HP-β-CD) for 3 days (100 mg kg⁻¹). On the following day, mice were anesthetized and eccentric injury of the gastrocnemius was performed using a 305B muscle lever system (Aurora Scientific Inc.)⁴. The knee was stabilized and the foot firmly affixed to a footplate on the torque sensor. Contraction was elicited by percutaneous electrical stimulation of the sciatic nerve. Optimal isometric tetanic torque was determined with increasing the current, with a minimum of 30 s between each stimulation to prevent fatigue. Eccentric contractions were created by translating the footplate 30° backward at a velocity of 40 mm s⁻¹ after the first 200 ms of the isometric contraction. Eccentric contractions at maximal isometric torque (0.2-ms pulse train at 100 Hz) were assayed 20 times with 1 min pauses between each, for the resistance to muscle damage. The decrease in the peak isometric force before the eccentric phase was used as a measure of damage. Following injury, the gastrocnemius was excised and flash frozen for western blotting.

In vivo manipulation of cardiac workload

Mice (9–11 months) were treated with either LC-1 or vehicle (HP-β-CD) for 3 days (100 mg kg⁻¹). On the subsequent day, mice were lightly anesthetized and instrumented for non-invasive ECG (Biopaq; three lead). ECG was monitored and digitized (1KHz) for 5 min before a challenge with isoproterenol (ISO; 0.5 mg kg⁻¹) and then for 45 min or until the heart rate was significantly depressed, at which time the mouse was euthanized under anaesthesia. Hearts were excised and flash frozen for western blotting.

Calcium stress assay

FDB muscle fibres from LC-1 and vehicle-treated mice were isolated as described above, placed into Hepes-buffered Ringer’s solution and plated into ECM-coated wells of a clear-bottom 96-well plate. Cells were loaded with Fluo-4 (5 μM) for 30 min and rinsed with Ringer’s solution. The cells of individual wells were electrically stimulated using a custom perfusion and stimulation apparatus (Four-Hour Day Foundation) for 5 min at 20 Hz, during which time Ca²⁺ was recorded.

Western blotting

For western blotting, 20 mg of clarified muscle or C2C12 extract was subjected to SDS-polyacrylamide gel electrophoresis, transferred on nitrocellulose membranes, and washed/blocked in 5% milk solution in PBS for 1 h. The membrane was probed overnight with primary antibody at room temperature. The primary antibodies were: anti-α-tubulin (Sigma DM1A, 1:1,000), anti-detyrosinated tubulin (AbCam #ab48389, 1:1,000), anti-acetylated tubulin (Sigma 6-11-B, 1:200), anti-Δ2-tubulin (Novus ##NB100-57397, 1:1,000), anti-polyglutamylation (Adipogene GT335, 1:1,000) and anti-gp91^phox (BD Transduction Labs #611415, 1:1,000). Membranes were washed two times for 10 min in 5% milk solution at room temperature and incubated with appropriate secondary antibody (1:10,000) for 1 h at room temperature, and the membranes were washed in 0.5% Tween solution in PBS two times for 10 min. Membranes were then exposed to enhanced chemiluminescence with SuperSignalWest Pico Chemiluminescent Substrate to develop immunoblots. Blots were imaged and quantified with an imaging system (Syngene, G:Box; GeneTools software), normalized to total protein (via PonceauS) or GAPDH (Fig. 1, cardiomyocytes) and analysed using ImageJ. Images were cropped for presentation. Uncropped scans of western blots are shown in Supplementary Fig. 6.

Statistical analysis

The sample number for each experiment was determined by a power analysis (http://www.power-analysis.com/) assuming a Cohen’s maximum variability of the population mean (determined by variance in published experiments) and a type 1 error of 0.5%. For in vivo studies, animals were randomly assigned to treatment groups and blinded for experimentation and analysis. All other experiments were performed and analysed unblinded. Normality of data sets was ensured for each statistical calculation. Significance was set a priori at P<0.05.

Additional information

How to cite this article: Kerr, J. P. et al. Detyrosinated microtubules modulate mechanotransduction in heart and skeletal muscle. Nat. Commun. 6:8526 doi: 10.1038/ncomms9526 (2015).

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Acknowledgements

Support was provided by the NIH (R01-AR062554 to CWW; R00 HL114879 to BLP; T32-AR07592 to J.P.K.; T32-AR053461 to PR), Hyundai Hope on Wheels (Hope Grant award to D.A.H.) and The V Foundation for Cancer Research (V Scholar award to D.A.H.). We thank Brunella Tedesco for her assistance with the AFM measurements. Assistance with the instrumentation and reagents for the myofibre stretch studies was kindly provided by IonOptix LLC and Aurora Scientific Inc. We thank E. Michael Ostap for careful review of the manuscript.

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Authors and Affiliations

Department of Orthopaedics, University of Maryland School of Medicine, Baltimore, 21201, Maryland, USA
Jaclyn P. Kerr, Guoli Shi & Christopher W. Ward
Department of Physiology, Pennsylvania Muscle Institute, Perelman School of Medicine, University of Pennsylvania, Philadelphia, 19104, Pennsylvania, USA
Patrick Robison, Alexey I. Bogush & Benjamin L. Prosser
Department of Medicinal Chemistry, College of Pharmacy, University of Minnesota, Minneapolis, 55455, Minnesota, USA
Aaron M. Kempema, Joseph K. Hexum & Daniel A. Harki
Department of Informatics, Bioengineering, Robotics and System Engineering, University of Genova, Genova, 16146, Italy
Natalia Becerra & Roberto Raiteri
Marlene and Stuart Greenebaum National Cancer Institute Cancer Center, University of Maryland School of Medicine, Baltimore, 21201, Maryland, USA
Stuart S. Martin
Center for Biomedical Engineering and Technology (BioMET), University of Maryland School of Medicine, Baltimore, 21201, Maryland, USA
Christopher W. Ward

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Jaclyn P. Kerr
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Patrick Robison
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Guoli Shi
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Alexey I. Bogush
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Roberto Raiteri
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Benjamin L. Prosser
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Christopher W. Ward
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Contributions

J.P.K., B.L.P., P.R., G.S., R.R., N.B. and C.W.W. performed experiments; A.M.K., J.K.H., S.S.M. and D.A.H. contributed reagents; J.P.K., B.L.P. and C.W.W. wrote the manuscript; all authors interpreted data and edited the manuscript.

Corresponding authors

Correspondence to Benjamin L. Prosser or Christopher W. Ward.

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B.L.P. and C.W.W. hold a US patent for MyoTak adhesive (patent number US 20120034620 A1), licensed to IonOptix LLC. C.W.W. holds a US patent for microtubule-targeted interventions for the muscular dystrophies (patent number US 2014015664 A1). The remaining authors declare no competing financial interests.

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Kerr, J., Robison, P., Shi, G. et al. Detyrosinated microtubules modulate mechanotransduction in heart and skeletal muscle. Nat Commun 6, 8526 (2015). https://doi.org/10.1038/ncomms9526

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Received: 03 February 2015
Accepted: 01 September 2015
Published: 08 October 2015
DOI: https://doi.org/10.1038/ncomms9526

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