Calpain-1 is required for hydrogen peroxide-induced myotube atrophy

Abstract

Recent reports suggest numerous roles for cysteine proteases in the progression of skeletal muscle atrophy due to disuse or disease. Nonetheless, a specific requirement for these proteases in the progression of skeletal muscle atrophy has not been demonstrated. Therefore, this investigation determined whether calpains or caspase-3 is required for oxidant-induced C2C12 myotube atrophy. We demonstrate that exposure to hydrogen peroxide (25 μM H₂O₂) induces myotube oxidative damage and atrophy, with no evidence of cell death. Twenty-four hours of exposure to H₂O₂ significantly reduced both myotube diameter and the abundance of numerous proteins, including myosin (−81%), α-actinin (−40%), desmin (−79%), talin (−37%), and troponin I (−80%). Myotube atrophy was also characterized by increased cleavage of the cysteine protease substrate αII-spectrin following 4 h and 24 h of H₂O₂ treatment. This degradation was blocked by administration of the protease inhibitor leupeptin (10 μM). Using small interfering RNA transfection of mature myotubes against the specific proteases calpain-1, calpain-2, and caspase-3, we demonstrated that calpain-1 is required for H₂O₂-induced myotube atrophy. Collectively, our data provide the first evidence for an absolute requirement for calpain-1 in the development of skeletal muscle myotube atrophy in response to oxidant-induced cellular stress.

skeletal muscle atrophy occurs as a consequence of numerous pathological conditions, including cachexia and disuse-related events (reviewed in Refs. 27 and 34). Numerous cellular events contribute to muscle atrophy, including those regulating protein synthesis, protease activation, ubiquitin conjugation, and autophagy (9, 17, 30, 31, 37, 39, 40). In recent years, progress has been made toward understanding the contributions of both ubiquitin ligase/proteasomal and lysosomal/autophagic-mediated skeletal muscle degradation processes. Although it has been argued that activation of the ubiquitin-proteasome system may play a dominant role in skeletal muscle atrophy (4, 21, 34), the role of cellular proteases in skeletal muscle atrophy is not well defined. In fact, new evidence suggests that a coordinated network of degradative processes is responsible for muscle atrophy (40).

Myofilament proteins make up the majority of protein in skeletal muscle cells, and the proteasome is important for the degradation of these proteins during disuse (reviewed in Ref. 4). Evidence reveals that the ubiquitin-proteasome system cannot degrade intact myofibril proteins, indicating that myofilament release is an initial and required step for proteasome-mediated degradation of myofibrillar proteins (4, 7, 20, 26, 27, 34). The inhibition of cysteine proteases (i.e., calpain and caspase-3) in skeletal muscle attenuates disuse muscle atrophy (16, 19, 38). We and others have hypothesized that disturbances in redox balance are critical regulatory mechanisms for the activation of both the Ca²⁺-dependent calpains I (μ-) and II (m-) and caspase-3 during atrophy and that these proteases are responsible for myofilament release during the initial stages of muscle atrophy (26, 27, 32).

Oxidative stress has been linked to skeletal muscle atrophy in numerous models of muscle wasting, including inactivity (27), diabetes (18), cancer (12), heart failure (13), chronic alcohol intake (25), human immunodeficiency virus infection (24), and sepsis (23). Although oxidative stress can activate cysteine proteases, the specific role that calpains and caspase-3 play in oxidant-induced muscle atrophy remains unclear. Therefore, this study investigated whether activation of cysteine proteases is a requirement for oxidant-induced skeletal muscle atrophy. Specifically, using both a pharmacological approach and RNA interference technology, we determined whether activation of calpain-1, calpain-2, or caspase-3 is required for hydrogen peroxide (H₂O₂)-induced skeletal muscle C2C12 myotube atrophy. We hypothesized that these cysteine proteases are required for H₂O₂-induced myotube atrophy in vitro. Our findings demonstrate a specific role for calpain-1 in H₂O₂-induced skeletal muscle atrophy in this model.

METHODS

Cultured myogenic cell line.

Myoblasts derived from mouse skeletal muscle (C2C12 cells; American Type Culture Collection, Rockville, MD) were cultured on six-well dishes in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% newborn calf serum, 1% penicillin/streptomycin, and 0.1% fungizone at 37°C in the presence of 5% CO₂ until 80% confluence was reached as visualized by light microscopy. Myotube differentiation was then initiated by replacement of the growth medium with differentiation medium: DMEM supplemented with 2% heat-inactivated horse serum, 1% penicillin/streptomycin, and 0.1% fungizone at 37°C in the presence of 5% CO₂, for 7 days. The muscle cells were examined for evidence of myotube formation and growth by light microscopy. H₂O₂ treatment of C2C12 cells involved dilution in differentiation media to final desired concentrations and standard incubation (37°C in the presence of 5% CO₂). H₂O₂-treated media were replaced at 12-h intervals for incubation times greater than 12 h.

Cell viability.

H₂O₂-induced alterations in C2C12 viability were analyzed via Trypan blue exclusion assay as previously described (36). Briefly, fully differentiated cells were treated with varying concentrations of H₂O₂ (0 μM control, 25 μM, 50 μM, 100 μM, and 200 μM) for predetermined durations (1 h, 2 h, 4 h, 24 h, 48 h, and 72 h). Media were removed, and the cells were incubated; 0.25% Trypan blue (Sigma) in PBS was applied to cultures for 5 min, and cultures were then rinsed twice with PBS. As a positive control, some untreated cultures were exposed to 80% ethanol in PBS for 30 min before application of Trypan blue. Images were obtained via an inverted microscope (Carl Zeiss Axiovert 200) camera system at ×10 magnification. Approximately 150 myotubes per treatment, a number chosen by determination of no additional change in standard deviation, were counted for the inclusion/exclusion of dye at each concentration and duration by a blinded investigator, and the percentage of viable cells (excluding dye) was determined.

Transient transfections.

Differentiated myotubes were transiently transfected in six-well plates with small interfering RNA (siRNA) sequences for cysteine proteases. At least three validated siRNA sequences (Ambion) were evaluated for knockdown efficacy for each cysteine protease. Efficiency of knockdown was deemed adequate when mRNA abundance for each protease was reduced by ∼70%. RNA oligos were transfected into differentiated myotubes using Lipofectamine 2000 (Invitrogen), according to the manufacturer's instructions. Briefly, siRNA and Lipofectamine 2000 were separately diluted in Opti-MEM (Gibco). The diluted Lipofectamine 2000 reagent was incubated for 5 min and was then added to the siRNA mixture at a final volume of 500 μl/well. The lipid/siRNA mixtures were allowed to complex for 20 min during which the cells were rinsed twice with Opti-MEM and bathed in a final volume of 2 ml Opti-MEM/well. Lipid:siRNA complexes were then applied to the cells (final transfection volume of 2.5 ml). Separate experiments were performed using equivalent amounts of lipid reagent devoid of siRNA. Cultures were rocked every 2 h for the initial 6 h of incubation and were incubated with the transfection mixture for a total of 24 h. After 24 h, transfection medium was removed and replaced with differentiation medium. Parallel cultures were transfected with fluorescein (FAM)-labeled nonsilencing control and calpain-1 siRNA oligomers (Ambion) and were imaged on an inverted fluorescence microscope (Carl Zeiss Axiovert 200) camera system at ×10 magnification to determine transfection efficiency. Cultures were maintained for an additional 24 h before H₂O₂ treatment.

Oxidative stress.

4-Hydroxynonenal (trans-4-hydroxy-2-nonenal; 4-HNE, C₉H₁₆O₂) is an α,β-unsaturated hydroxyalkenal that is produced by lipid peroxidation in cells (35). 4-HNE modification of proteins was analyzed as an indicator of oxidative damage in C2C12 cells via Western blotting as described (1). Values for 4-HNE were corrected for duration-matched untreated controls (0 μM H₂O₂) and are presented as fold changes.

Myotube atrophy.

Myotube diameter morphological analysis was performed as previously described (22) with the following modifications. Briefly, images of cultures were obtained via phase contrast at ×100 magnification after the appropriate treatment durations were obtained on an inverted microscope (Carl Zeiss Axiovert 200) camera system at ×10 magnification. The diameters were measured in a total of 100 myotubes from at least 10 random fields, a number chosen by determination of no additional change in standard deviation, using computerized image analysis (Scion Image, Frederick, MD). Each myotube analyzed was measured at three points along the length of the myotube in a blinded fashion, and results are expressed as percentage of the control treatment diameter.

Western blot analysis.

On completion of the appropriate incubation time, cells were rinsed 2× in ice-cold 1× PBS and scraped for protein isolation in 130 μl nondenaturing lysis buffer (NDL buffer: 1% Triton X-100, 300 mM NaCl, 50 mM Tris-base, 5 mM EDTA, 3.1 mM sodium azide, 95 mM NaF, 22 μM Na₃VO₄), vortexed, incubated at 4°C for 25 min, and centrifuged at 1,000 g for 5 min. The supernatants were subsequently assayed for protein using the Bradford method (Sigma, St. Louis, MO), separated by polyacrylamide gel electrophoresis via 4–15% gradient, and transferred to nitrocellulose membranes (100 V for 3 h at 4°C) for Western blot analysis. As verification of equal loading and transfer, the resulting transfer membrane was stained with Ponceau S, and each lane was analyzed using computerized image analysis and used to correct the Western blot analysis for the total amount of protein in each lane, respectively (Scion Image) (data not shown). Membranes were then probed for manganese superoxide dismutase (MnSOD; SOD-111; Stressgen; Victoria, BC, Canada), copper zinc superoxide dismutase (CuZnSOD; SOD-101; Stressgen), catalase (Ab16731; Abcam, Cambridge, MA) and calpain-1 (no. 2556), calpain-2 (no. 2539), caspase-3 (no. 9665), αII-spectrin (no. 2122), cleaved αII-spectrin (no. 2121), talin (no. 4021), and troponin I (no. 4002) (all purchased from Cell Signaling Technology, Carlsbad, CA). Myosin (A4335) and α-actinin (A7811) were purchased from Sigma-Aldrich (St. Louis, MO). Actin (sc-7210) and desmin (sc-14026) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). 4-Hydroxynonenal (trans-4-hydroxy-2-nonenal; 4-HNE, C₉H₁₆O₂; ab46545) was purchased from Abcam.

RNA isolation and cDNA synthesis.

Cells harvested for RNA isolation were rinsed two times in ice-cold 1× PBS and scraped in TRIzol reagent (Life Technologies, Carlsbad, CA), according to the manufacturer's instructions. Total RNA (5 μg) was then reverse transcribed using the Superscript III First-Strand Synthesis System for RT-PCR (Life Technologies) using oligo(dT)20 primers and the protocol outlined by the manufacturer.

Real-time polymerase chain reaction.

One microliter of cDNA was added to a 25-μl PCR reaction for real-time PCR using Taqman chemistry and the ABI Prism 7000 Sequence Detection System (ABI, Foster City, CA). Relative quantitation of gene expression in treated C₂C₁₂ cells was performed using the comparative computed tomography method (ABI, User Bulletin no. 2). This method uses a single sample, the calibrator sample (hprt; GenBank NM_013556.2), for comparison of every unknown sample's gene expression. ΔΔC_T [ΔC_T (calibrator) − ΔC_T (sample)] was then calculated for each sample, and relative quantification was calculated as 2ΔΔC_T (where C_T is cycle threshold). Hypoxanthine-guanine phosphoribosyltransferase (Hprt), an enzyme involved in purine catabolism, was chosen as the reference gene on the basis of initial experiments showing unchanged expression with our experimental manipulations. Fivefold dilution curves were assayed on selected samples to confirm the validity of this quantitation method for each gene. Calpain-1 (GenBank NM_001110504.1), calpain-2 (GenBank NM_009794.3), caspase-3 (GenBank NM_009810.2), and calpastatin (GenBank NM_009817.3) mRNA transcripts were assayed using predesigned rat primer and probe sequences commercially available from Applied Biosystems (Assays-on-Demand).

Statistical analysis.

A priori analysis was used to determine any differences between control treatments of varying durations. When no differences existed between groups, respective groups were pooled. Comparisons between treatments were made by a one-way analysis of variance and, when appropriate, a Tukey honestly significant difference test was performed post hoc. Significance was established at P < 0.05. Values are reported as means ± SE.

RESULTS

Cell viability and oxidative stress.

The toxicity of H₂O₂ treatment to differentiated C2C12 myotubes was determined using the Trypan blue exclusion assay in an attempt to determine the appropriate concentration of peroxide and duration of administration for our experiments. Only H₂O₂ concentrations of 25 μM or 50 μM failed to induce myotube viability loss at all durations of exposure (1 h, 2 h, 4 h, 24 h, 48 h, or 72 h; Fig. 1A). We then tested these nontoxic H₂O₂ concentrations to determine whether they were sufficient to induce myotube oxidative damage. H₂O₂ concentration of 25 μM or 50 μM, regardless of the duration of administration (4 h, 24 h, 48 h, or 72 h), resulted in increased 4-HNE modification of myotube proteins (Fig. 1B and Supplemental Fig. S1A; supplemental material for this article is available online at the American Journal of Physiology: Cell Physiology website).

Fig. 1.

Cell viability and oxidative stress in C2C12 myotubes treated with H₂O₂. A: cell viability. Fully differentiated myotubes were treated with graded concentrations (0 μM vehicle, 25 μM, 50 μM, 100 μM, or 200 μM final concentrations of H₂O₂) for varying durations (1 h, 2 h, 4 h, 24 h, 48 h, or 72 h). Cells were stained with Trypan blue and analyzed for the percentage of total cells excluding dye (cell viability %). Values are representative of 6 independent experiments and presented as means ± SE. *Significantly different from duration matched 0 μM (control) concentration. B: oxidative stress. Fold induction (Ind) α,β-unsaturated hydroxyalkenal [4-hydroxynonenal (trans-4-hydroxy-2-nonenal) 4-HNE, C₉H₁₆O₂; lipid peroxidation by-product] modification of proteins. On the basis of cell viability findings, fully differentiated myotubes were treated with graded concentrations (0 μM vehicle, 25 μM, or 50 μM final concentrations of H₂O₂) for varying durations (4 h, 24 h, 48 h, or 72 h). Values are representative of 6 independent experiments, normalized to duration-matched 0 μM (control) concentration of H₂O₂, and presented as means ± SE. *Significantly (P < 0.05) different from duration-matched 0 μM (control).

Antioxidant enzyme expression.

We also analyzed the protein expression of catalase, CuZnSOD, and MnSOD to determine whether the administration of 25 μM H₂O₂ alters the baseline abundance of endogenous myotube antioxidants (Supplemental Fig. S1B). There were no alterations in the abundance of any antioxidant enzyme analyzed after 24 h of 25 μM H₂O₂ treatment. After extended periods of exposure, however, CuZnSOD optical density (OD) (48 h: 1.88 ± 0.11 OD; 72 h: 4.24 ± 0.59 OD) and MnSOD (72 h: 1.51 ± 0.10 OD) protein abundances increased from control values (1.0 ± 0.07 OD and 1.0 ± 0.05 OD, respectively).

H₂O₂-induced myotube atrophy.

Myotube diameter was analyzed as an indicator of myotube atrophy in response to 25 μM H₂O₂ treatment. Myotube diameter was not altered from control values following 4 h of peroxide treatment (91.78 ± 7.9% of control). Myotube diameter decreased 31–37% with 25 μM H₂O₂ treatment during 24, 48, or 72 h. This finding demonstrated 2 important points: 1) that this lower dose of H₂O₂ treatment was sufficient to induce myotube atrophy and 2) that the 24-h duration of exposure resulted in roughly the same level of myotube atrophy as the 72 h.

Sarcomeric protein expression.

Four hours of 25 μM H₂O₂ exposure resulted in decreases in myosin (−22%), α-actinin (−23%), and desmin (−80%) protein expressions (Table 1 and Supplemental Fig. S1C). Interestingly, 4 h of exposure also increased (101%) the protein expression of troponin I. Twenty-four hours of 25 μM H₂O₂ exposure resulted in further decreases in myosin and desmin, and decreases in talin (−37%), troponin I (−80%), and αII-spectrin (−29%) proteins. Actin protein expression was not altered by H₂O₂ exposure at any time-point analyzed. Overall, these data suggest that H₂O₂ exposure differentially alters the expression of numerous sarcomeric proteins in atrophying myotubes.

Table 1.

Sarcomeric protein abundance

	Duration of 25 μM H2O2 Treatment
	Con	4 h	24 h
Myosin	1.00±0.11	0.78±0.06*	0.19±0.05*†
Actin	1.00±0.03	1.02±0.08	1.12±0.03
α-Actinin	1.00±0.05	0.77±0.07*	0.60±0.01*†
Desmin	1.00±0.18	0.20±0.01*	0.21±0.01*
Talin	1.00±0.05	0.96±0.09	0.63±0.03*†
Troponin I	1.00±0.29	2.01±0.20*	0.20±0.07*†
αII-Spectrin	1.00±0.08	1.26±0.27	0.71±0.06*†

Values are means ± SE and are representative of 6 independent experiments. Sarcomeric protein abundance decreases in C2C12 myotubes with 25 μM H₂O₂ treatment. Protein expression was verified by Western blot analysis.

^*Significantly different (P < 0.05) from control (Con).

^†Significantly different (P < 0.05) from 4-h treatment duration.

Cysteine protease requirement for H₂O₂-induced myotube atrophy.

A key role for calpain and caspase cysteine proteases in the regulation of skeletal muscle size is their cleavage of sarcomeric protein substrates (including αII-spectrin), resulting in the disorganization of normal sarcomeric architecture (4, 29). The percentage of total αII-spectrin protein cleaved by proteases with 4 h or 24 h of 25 μM H₂O₂ treatment increased (Fig. 2A), indicating the amount of relative alterations in cleaved αII-spectrin and total αII-spectrin with H₂O₂ treatment. Furthermore, addition of the nonspecific protease inhibitor leupeptin to the differentiation medium attenuated myotube diameter loss associated with 24 h of 25 μM H₂O₂ treatment (Fig. 2B).

Fig. 2.

Protease activity in H₂O₂ induced atrophying myotubes. A: αII-spectrin protease substrate cleavage. αII-spectrin fragments are representative of the calpain-generated 150- and 145-kDa fragments. Fully differentiated C2C12 myotubes were treated with 0 μM vehicle (control) or 25 μM H₂O₂ for 4 h or 24 h. Cellular lysates were collected and analyzed via Western blot for the sarcomeric protease substrate αII-spectrin. Values are representative of 6 independent experiments and are presented as the % cleaved αII-spectrin protein, means ± SE. *Significantly (P < 0.05) different from control. B: myotube diameters. Fully differentiated C2C12 myotubes were treated with 0 μM vehicle (control), 25 μM H₂O₂ for 24 h, or 25 μM H₂O₂ and 10 μM leupeptin (broad spectrum protease inhibitor; Leup) for 24 h. Representative images were acquired and analyzed for myotube diameters. Values are representative of 6 independent experiments and are presented as the percentage of 0 μM vehicle control values, means ± SE. *Significantly (P < 0.05) different from control. †Significantly (P < 0.05) different from 25 μM H₂O₂.

To determine the specific cysteine protease (i.e., calpain or caspase-3) required for the development of myotube atrophy, we tested the efficiency and specificity of calpain-1, calpain-2, and caspase-3 RNA interference (Table 2). After the establishment of specific and efficient knockdown of only the mRNA abundances and protein expressions of the targeted proteases, myotubes were transfected with each siRNA individually, subsequently treated with 25 μM H₂O₂ for 24 h, and diameter measurements were obtained (Fig. 3A). Calpain-1 knockdown protected C2C12 myotubes from diameter loss during peroxide treatment (Fig. 3B). Calpain-2 or caspase-3 interference proved ineffective in protecting the myotubes from atrophy (Fig. 3B).

Table 2.

Efficiency and specificity of protease siRNA in myotubes

	Treatment Group
	Calp-1 siRNA	Calp-2 siRNA	Casp-3 siRNA
Protein expression
Calpain-1	0.31±0.04*	1.19±0.15	1.19±0.15
Calpain-2	0.93±0.05	0.41±0.05*	0.89±0.06
Caspase-3	0.78±0.10	0.84±0.11	0.38±0.05*
mRNA abundance
Calpain-1	0.29±0.10*	0.83±0.26	1.23±0.13
Calpain-2	1.07±0.10	0.31±0.07*	0.81±0.17
Caspase-3	1.33±0.16	1.21±0.29	0.28±0.04*

Values are means ± SE and are representative of 6 independent experiments. Efficiency and specificity of protease small interfering (si)RNA in C2C12 myotubes. C2C12 cells were allowed to differentiate for 4 days and were transfected with siRNA against calpain-1, calpain-2, or caspase-3. Cells were then treated and harvested for protein or RNA. Protein abundance was verified by Western blot analysis, and values are presented as fold induction from control siRNA-treated cells. mRNA expression was verified by real-time RT-PCR, and values are presented as corrected for hprt and are normalized to control siRNA-treated cells.

^*Significantly different (P < 0.05) from respective control cells.

Fig. 3.

Calpain-1 interference attenuates myotube atrophy. Fully differentiated C2C12 myotubes were transfected with control (mismatched), calpain-1, calpain-2, or caspase-3 small interfering (si)RNA and subsequently treated with 0 μM vehicle (control), or 25 μM H₂O₂ for 24 h. Representative images were acquired and analyzed for myotube diameters. A: representative myotube images of 0 μM vehicle (control), 25 μM H₂O₂, and 25 μM H₂O₂ + calpain-1 siRNA. B: myotube diameters. Values are representative of 6 independent experiments and are presented as the percentage of 0 μM vehicle controls values, means ± SE. *Significantly (P < 0.05) different from control. †Significantly (P < 0.05) different from 25 μM H₂O₂. ‡Significantly (P < 0.05) different from 25 μM H₂O₂ + calpain-1 siRNA.

The mRNA abundance for calpain-1 is transiently increased with 4 h of H₂O₂ treatment and returned to baseline with 12 h and 24 h of H₂O₂ treatment (Fig. 4A). Transfection with siRNA reduces mRNA abundance of calpain-1 independent of H₂O₂ administration. The mRNA abundance of the endogenous calpain inhibitor calpastatin increased only after 24 h of H₂O₂ treatment (Fig. 4B). Further analysis of the calpain-1 knockdown demonstrated a significant reduction in the early (4 h) autoproteolytic cleavage and activation of calpain protein (Fig. 5, A and B) and cleavage of αII-spectrin (Fig. 5, A and C). Neither calpain-2 nor caspase-3 interference reduced the H₂O₂ cleavage of αII-spectrin.

Fig. 4.

Time course of calpain-1 mRNA expression in H₂O₂-treated myotubes. Fully differentiated C2C12 myotubes were transfected with control (mismatched) or calpain-1 siRNA and subsequently treated with 0 μM vehicle (control), or 25 μM H₂O₂ for 4, 12, or 24 h. mRNA expression was determined by real-time RT-PCR and are presented as corrected for hprt and normalized to control siRNA-treated cells. A: calpain-1 mRNA. B: calpastatin mRNA. Values are means ± SE and are representative of 6 independent experiments. *Significantly different (P < 0.05) from respective control cells. †Significantly (P < 0.05) different from 25 μM H₂O₂.

Fig. 5.

Calpain-1 interference inhibits H₂O₂-induced protease activity. Fully differentiated C2C12 myotubes were transfected with control (mismatched) or calpain-1 siRNA and subsequently treated with either 0 μM vehicle (control) or 25 μM H₂O₂ for 4 h. Cellular lysates were collected and analyzed via Western blot for the sarcomeric protease substrate αII-spectrin as well as cleaved (activated) calpain-1. A: representative blot images of both total and cleaved calpain-1 and αII-spectrin. B: fold induction of autoproteolytically cleaved calpain-1 protein. Values are representative of 6 independent experiments, corrected for Ponceau-stained total protein within lane, normalized to control values, and presented as means ± SE. C: % αII-spectrin cleavage. Values are representative of 6 independent experiments, corrected for Ponceau-stained total protein within lane, and presented as the % cleaved αII-spectrin protein, means ± SE. *Significantly (P < 0.05) different from control. †Significantly (P < 0.05) different from 25 μM H₂O₂.

DISCUSSION

Overview of principle findings.

Several new and important findings emerged from these experiments. First, these results demonstrate that exposure of C2C12 myotubes to low levels of H₂O₂ is sufficient to induce myotube oxidative stress and atrophy within 24 h. This H₂O₂-induced atrophy is characterized by an early and sustained loss of numerous sarcomeric proteins and cleavage of the structural myofilament protein, αII-spectrin. Second, activation of the cysteine protease calpain-1 is a requirement for H₂O₂-induced atrophy. A brief discussion of these key results follows.

Requirement for calpain-1 in H₂O₂-induced myotube atrophy.

The most important finding from the current study is that the knockdown of calpain-1 attenuated H₂O₂-induced myotube atrophy. This suggests a direct requirement for calpain-1 in the progression of H₂O₂-induced myotube atrophy. The magnitude of protease knockdown in the current study was not sufficient to completely abrogate calpain mRNA abundance or protein expression. It is possible that residual levels of protease may continue to mediate the normal response to physiologically relevant stimuli, resulting in the attenuation but not complete prevention of atrophy or substrate cleavage. Our data clearly demonstrate that this level of interference was sufficient to protect the myotubes from atrophy and the cleavage of protease substrate proteins. The transfection efficiency observed in the current study combined with the large number of myotubes and sample sizes used for the analysis of diameter and biochemistries, however, suggest that residual physiologically activated calpains are insufficient to overcome the magnitude of protective effects of the protease knockdown achieved. The C2C12 myotubes used in the present study were analyzed after a treatment period encompassing a total of 7 days of differentiation, allowing for sufficient time for mature myotube formation. However, residual calpain may also be localized to nondifferentiated myoblasts or in nontransfected myotubes in culture, providing another potential mechanism for continued protease activation, mRNA abundance, and protein expression by analysis.

To date, the mechanistic link between oxidants and activation of calpain-1 remains unclear. A potential mechanism for calpain activation by reactive oxygen species (ROS) in skeletal muscle involves the formation of reactive aldehydes that damage calcium handling proteins and increase cytosolic Ca²⁺ levels. Our finding that exposure of myotubes to H₂O₂ increases cellular levels of the reactive aldehyde (4-HNE) is consistent with this notion. Regardless of the mechanism of how H₂O₂ activates calpain in myotubes, our data suggest that calpain-1 activity participates in myofilament protein release in atrophying muscles. Finally, while active calpain-1 participates in myofilament release, it is also possible that calpains could contribute to skeletal muscle atrophy in other ways as well. For example, additional reported functions of calpain proteases in skeletal muscle include transcription factor degradation and modification of protein kinase B (PKB/Akt) activity (reviewed in Ref. 34).

H₂O₂-induced muscle atrophy.

Oxidants such as H₂O₂ are theorized to play important roles in the progression of skeletal muscle atrophy due to inactivity or various diseases. In fact, antioxidant administration attenuates myofiber atrophy in both disease and disuse models (20, 25, 32). Hydrogen peroxide is often used as an exogenous oxidant treatment to promote oxidative stress and proteolytic signaling pathways in both isolated skeletal muscles (6, 28) and in vitro myotubes (14, 15). Although the ability of H₂O₂ to induce skeletal muscle myofiber atrophy has been inferred in several studies, the current study is the first to demonstrate a direct link between H₂O₂ and myotube atrophy. Moreover, our results also extend previous work (15) by demonstrating that proteases involved in the ubiquitin-proteasome proteolytic cascade are activated and transcriptionally induced by the treatment of C2C12 myotubes with H₂O₂. Note that we cannot attribute the effects of the H₂O₂ treatment solely to the biochemical properties of this oxidant. Indeed, it is possible that H₂O₂ could react with transition metals and/or superoxide in the cell to produce more reactive species such as the hydroxyl radical. Hence, the specific oxidant(s) directly responsible for our results remain unknown.

Recently, the classical dogma on oxidants as cellular disruptors of homeostasis by overwhelming endogenous antioxidant systems has undergone revision. A role for ROS in compartmentalized cellular signaling regulating cellular functions has been introduced (8, 10). Specifically, oxidants have been recognized as modifiers of cellular signal transduction, phosphatase and protease activity, chaperone proteins, and transcription factors (reviewed in Ref. 8). H₂O₂ is receiving amplified attention as a cellular signaling molecule due to its ability to reversibly oxidize amino acids, specifically cysteines (8), resulting in posttranslational protein modifications that affect cellular function. A continuum of redox-based cellular modifications, proposed by Forrester and Stamler (3), suggests that reversible posttranslational modifications of cysteine thiol side chains (S-nitrosothiol, sulfenic acid, and disulfides) are demonstrative of physiological signaling modifications initiated by oxidants. In contrast, irreversible modifications (protein carbonyl and methionine sulfoxide formation) are maladaptive and represent oxidative stress-induced cellular injury. In this context, the low dose of H₂O₂ used in the current study was insufficient to induce cell death but resulted in the induction of endogenous antioxidant (CuZnSOD and MnSOD) protein abundance, development of oxidative stress (4-HNE modification of proteins), and atrophy. It remains plausible, therefore, that H₂O₂ functions in multiple stages along the redox continuum, first as a signaling intermediate to initiate protein modifications and cellular signaling and later as promoter of oxidative injury. At present, the relative contributions of the overlapping phases of this redox continuum to the progression of skeletal muscle atrophy remains unknown.

Cysteine protease-mediated muscle proteolysis.

The involvement of cysteine proteases in skeletal muscle atrophy has been implicated in reports suggesting that inhibition of calpain or caspase-3 retards both disuse and sepsis-induced muscle atrophy (2, 16, 19, 38). We and others have postulated that myofilament protein release may be a rate-limiting step in myofilament protein degradation by the ubiquitin-proteasome system in atrophying skeletal muscle (7, 11, 21, 34). The current study further corroborates this idea by demonstrating that protease activation and myofilament substrate cleavage occur early during the development of atrophy in myotubes. In direct contrast to this theory, Zhao et al. (40) treated cells with both lysosomal and proteasomal inhibitors and demonstrated a 90% reduction in overall proteolysis. These investigators failed to inhibit overall muscle proteolysis with specific pharmacological inhibitors of caspases and concluded that caspases and/or calpains play a minor role in overall cellular proteolysis. The current findings corroborate a negligible role for caspase-3 in skeletal muscle atrophy due to H₂O₂ treatment. However, our results clearly demonstrate an important role for calpain-1 in oxidant-induced myotube atrophy. We appreciate that inhibition of the proteasome (4) would attenuate a portion of skeletal muscle proteolysis in response to atrophic stimuli. Nonetheless, since the proteasome cannot degrade intact myofilaments, it is feasible that proteolytic release of these proteins remains an initial and required step for the subsequent degradation of myofilament proteins via the proteasome proteolytic system. In support of this postulate, a recent report indicates that calpain inhibition can retard muscle proteolysis independent of key ubiquitin ligases deemed critical to the proteasome system (2). Therefore, we conclude that an important role for calpains in skeletal muscle atrophy remains.

Calpains are classically defined as calcium-activated proteases found in most cell types (5). Despite the fact that H₂O₂-induced atrophy and cellular oxidative stress result in calpain activation and myotube atrophy, a direct link between calcium release and oxidative stress has not been definitively proven in the current study. A possible link between cellular oxidative stress and cellular calcium overload involves the oxidant-induced production of reactive aldehydes (4-HNE) that inhibit plasma membrane Ca²⁺-ATPase activity and attenuate the ability of the cell to remove cytosolic Ca²⁺ (33). Our study demonstrates that the formation of reactive aldehydes is an initial event in response to the stress provided by H₂O₂ treatment of myotubes in vitro. This may provide insight into a possible mechanism for Ca²⁺-induced calpain activation in the current model. Future studies are needed to examine the role of oxidant stress in Ca²⁺-induced calpain activation and should investigate stress-induced alterations in Ca²⁺-ATPase membrane channel function or structure as a potential mechanism.

Conclusions.

Our study revealed that the development of hydrogen peroxide-induced myotube atrophy in culture requires calpain-1, but not calpain-2 or caspase-3 cysteine proteases. The exact role for calpain-1 activity in atrophying myotubes is not entirely clear, but it may be due in part to its cleavage of structural myofilament proteins in response to hydrogen peroxide treatment. Although a requirement for calpain-1 for all types of muscle atrophy has not been established, our discovery that calpain-1 is required for H₂O₂-induced atrophy could lead to refined pharmacological or genetic therapies targeted at the maintenance of protein balance in skeletal muscle during wasting conditions involving oxidative stress.

GRANTS

This work was supported by National Heart, Lung, and Blood Institute Grant RO1-HL-072789 (to S. K. Powers).