- Email: [email protected]
The effects of acetaldehyde and acrolein on muscle catabolism in C2 myotubes
Free Radical Biology and Medicine 65 (2013) 190–200
Contents lists available at SciVerse ScienceDirect
Free Radical Biology and Medicine journal homepage: www.elsevier.com/locate/freeradbiomed
Original Contribution
The effects of acetaldehyde and acrolein on muscle catabolism in C2 myotubes Oren Rom a, Sharon Kaisari a, Dror Aizenbud a,b, Abraham Z. Reznick a,n a b
Department of Anatomy and Cell Biology, Rappaport Faculty of Medicine, Technion–Israel Institute of Technology, Bat Galim, Haifa 31096, Israel Orthodontic and Craniofacial Department, Rambam Health Care Campus, Bat Galim, Haifa 31096, Israel
art ic l e i nf o
a b s t r a c t
Article history: Received 24 February 2013 Received in revised form 23 May 2013 Accepted 12 June 2013 Available online 20 June 2013
The toxic aldehydes acetaldehyde and acrolein were previously suggested to damage skeletal muscle. Several conditions in which exposure to acetaldehyde and acrolein is increased were associated with muscle wasting and dysfunction. These include alcoholic myopathy, renal failure, oxidative stress, and inflammation. A main exogenous source of both acetaldehyde and acrolein is cigarette smoking, which was previously associated with increased muscle catabolism. Recently, we have shown that exposure of skeletal myotubes to cigarette smoke stimulated muscle catabolism via increased oxidative stress, activation of p38 MAPK, and upregulation of muscle-specific E3 ubiquitin ligases. In this study, we aimed to investigate the effects of acetaldehyde and acrolein on catabolism of skeletal muscle. Skeletal myotubes differentiated from the C2 myoblast cell line were exposed to acetaldehyde or acrolein and their effects on signaling pathways related to muscle catabolism were studied. Exposure of myotubes to acetaldehyde did not promote muscle catabolism. However, exposure to acrolein caused increased generation of free radicals, activation of p38 MAPK, upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1, degradation of myosin heavy chain, and atrophy of myotubes. Inhibition of p38 MAPK by SB203580 abolished acrolein-induced muscle catabolism. Our findings demonstrate that acrolein but not acetaldehyde activates a signaling cascade resulting in muscle catabolism in skeletal myotubes. Although within the limitations of an in vitro study, these findings indicate that acrolein may promote muscle wasting in conditions of increased exposure to this aldehyde. & 2013 Elsevier Inc. All rights reserved.
Keywords: Acetaldehyde Acrolein Muscle catabolism Oxidative stress p38 MAPK E3 ubiquitin ligases Free radicals
Aldehydes are highly reactive and toxic molecules. Exogenous exposure to aldehydes has been associated with increased risk of various pathologies and death [1]. Aldehydes are also produced endogenously during metabolism of amino acids and lipids and are involved in numerous biochemical and physiological processes. Acetaldehyde and acrolein are toxic aldehydes and their involvement in several pathologies has been studied extensively in the past [2]. The saturated aldehyde acetaldehyde is mainly produced by metabolism of alcohol in the liver and is considered to be responsible for the pathological complications of alcohol consumption [2]. Acetaldehyde has been previously suggested to damage skeletal muscle in alcoholism [3,4]. Approximately 50% of alcohol misusers suffer from chronic alcoholic myopathy in
Abbreviations: CS, cigarette smoke; MyHC, myosin heavy chain; MAPK, mitogenactivated protein kinase; atrogin-1, muscle atrophy F-box protein; MuRF1, muscle ring finger-1 protein; NAC, N-acetylcysteine; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; ROS, reactive oxygen species; RNS, reactive nitrogen species; GSH, reduced glutathione n Corresponding author. Fax: +97248295403. E-mail address:[email protected] (A.Z. Reznick). 0891-5849/$ - see front matter & 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.freeradbiomed.2013.06.024
which muscle mass may be reduced by up to 30%. In alcoholics, blood levels of acetaldehyde are relatively high and acetaldehyde was suggested to be a primary factor in alcohol-induced muscle damage [4]. Previous studies showed that both in vitro exposure of myocytes to acetaldehyde and injection of acetaldehyde to rats resulted in low synthesis of muscle proteins [5,6]. Acrolein is an extremely toxic and highly reactive α,β-unsaturated aldehyde that has been linked with various diseases and shown to regulate activation of transcription factors and signaling molecules [7]. Endogenous sources of acrolein include lipid peroxidation, in which highly reactive aldehydes are produced [2,8]. These aldehydes have been recognized as causative factors in aging and age-associated diseases, including cardiovascular disease and diabetes [8]. Acrolein can also be produced in situations of oxidative stress and inflammation during degradation of threonine by myeloperoxidase and degradation of spermine and spermidine by amine oxidase [9]. High serum or tissue levels of acrolein have been described in various pathological conditions such as renal failure, diabetes, and oxidative stress [10–12]. Wasting and dysfunction of skeletal muscle are common in both renal failure and diabetic patients [13]. Oxidative stress and chronic inflammation in which acrolein levels may be increased
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
are known to activate signaling pathways that lead to atrophy and loss of skeletal muscle fibers [9,14]. Thus, high levels of acrolein in these pathological conditions may exacerbate the catabolism of skeletal muscle. Previous studies have investigated the effects of acrolein exposure on cardiac and smooth muscle. In rodents, acrolein exposure was shown to induce cardiomyopathy characterized by increased oxidative stress and upregulation of proinflammatory cytokines [1,15]. In smooth muscle cultures treated with acrolein, activation of various signaling pathways, including the mitogen-activated protein kinase (MAPK) pathway, was demonstrated [16–18]. However, as far as we are aware, the effects of acrolein on skeletal muscle have not been examined previously. Cigarette smoke (CS) is a major exogenous source of both acetaldehyde and acrolein [2,7,9]. CS from a common cigarette contains approximately 1.5 mg of acetaldehyde. Non-proteinbound acetaldehyde can cross the alveolar–capillary membranes of the lungs and cigarette smoking can increase blood concentrations of acetaldehyde [19]. CS contains 60–100 mg of acrolein per cigarette [20]. Saliva, blood, and urine levels of acrolein and its metabolites are increased in smokers in comparison with nonsmokers [10,21–24]. Previous studies have revealed an association between smoking and skeletal muscle damage. Increased muscular damage and expression of genes associated with muscle catabolism were found in skeletal muscle biopsies taken from smokers and compared with nonsmokers [25,26]. In addition, smoking has been previously associated with sarcopenia, the age-related loss of muscle mass and strength [27–29]. Recently, we have shown that CS exposure to skeletal myotubes caused cell atrophy and degradation of myosin heavy chain (MyHC) via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ligases of the ubiquitin–proteasome system: muscle atrophy F-box protein (atrogin-1) and muscle ring finger-1 protein (MuRF1) [30]. Acetaldehyde and acrolein present in CS were previously suggested to possess potential catabolic effects on skeletal muscle [31]. Increased levels of acetaldehyde and acrolein are evident in various states associated with muscle catabolism, including oxidative stress, inflammation, renal failure [9–12], and alcoholism [2–4]. The aim of this study was to investigate the effects of acetaldehyde and acrolein on catabolism of skeletal myotubes. This was done by exposing C2 skeletal myotubes to various levels acetaldehyde and acrolein. We hypothesized that in a manner similar to that of CS, exposure of cultured myotubes to acetaldehyde and acrolein will lead to muscle atrophy and degradation of muscle proteins via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ubiquitin ligases atrogin-1 and MuRF1.
Materials and methods Cell culture The C2 cell line of mouse skeletal myoblasts was a generous gift from Professor Eyal Bengal (Rappaport Faculty of Medicine, Technion, Israel). C2 myoblasts were grown as previously described [30] in 24-well, 35-mm, and 100-mm plates in growth medium (GM) consisting of Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% (v/v) heat-inactivated fetal bovine serum, 1% (v/v) penicillin/streptomycin, and 1% (v/v) L-glutamine at 37 1C in humidified 95% air:5% CO2 atmosphere. For differentiation of myotubes, myoblasts were plated in 0.1% gelatin-coated plates and were grown to 90% confluence. At this point GM was replaced by differentiation medium (DM) consisting of DMEM supplemented with 2% (v/v) heat-inactivated horse serum, 1% (v/v) penicillin/ streptomycin, and 1% (v/v) L-glutamine. During differentiation, DM
191
was replaced every 48 h for 6 days until cell fusion and formation of multinucleated myotubes were achieved. Successful cell differentiation was determined by expression of the main contractile protein MyHC as measured by immunoblotting. Cell media and chemicals were purchased from Biological Industries (Bet HaEmek, Israel). Cell treatments Experiments were conducted on day 7 of differentiation when the cells have completed their differentiation into elongated multinucleated myotubes. Myotubes were exposed to various levels of acetaldehyde or acrolein (Sigma–Aldrich, St. Louis, MO, USA) freshly prepared in DM to final concentrations of 10–500 mM. In each experiment, medium was replaced by fresh DM containing acetaldehyde or acrolein. Control myotubes were treated with fresh DM without acetaldehyde or acrolein. Myotubes were then incubated at 37 1C for various time intervals. In some experiments, myotubes were pretreated with the antioxidant N-acetylcysteine (NAC) or with SB203580, a specific inhibitor of p38 MAPK (Sigma– Aldrich). Myotubes were treated with NAC (2 mM) or SB203580 (5 mM) prepared in fresh DM and incubated for 1 h or 15 min, respectively. Then, medium was replaced by DM containing acetaldehyde or acrolein and myotubes were transferred for incubation at 37 1C. NAC at concentrations of 0.1–30 mM was previously demonstrated to have antioxidant properties in various in vitro systems [32]. Clinically, peak plasma concentration of NAC reached 3.4 mM after intravenous administration of NAC at 150 mg/kg [32]. Estimation of cell viability To assess the effects of acetaldehyde and acrolein on viability of myotubes, 2 105 cells were seeded in 24-well plates and grown to 90% confluence for differentiation to myotubes. On day 7 of differentiation, myotubes were treated with DM containing acetaldehyde or acrolein and incubated for increasing time points (1, 2, 6, 24 h). After treatment, the viability of myotubes was determined by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay (5 mg/ml; Sigma–Aldrich) as previously described [30]. The OD was spectrophotometrically measured in triplicate for each experiment at 570 nm using an ELISA reader (Biochrom Anthos Zenyth, Cambridge, UK). Viability was expressed as a percentage of the values of control myotubes (corresponding to 100%). Estimation of ROS generation To assess the effects of acetaldehyde and acrolein on generation of ROS, 5 105 cells were seeded in 35-mm plates and grown to 90% confluence for differentiation to myotubes. On day 7 of differentiation, myotubes were treated with acetaldehyde or acrolein. After treatment, ROS generation was measured by 2′,7′dichlorofluorescein (DCF) assay as previously described [30]. Briefly, DNA of nuclei was stained with 1.5 mg/ml Hoechst (Invitrogen, Carlsbad, CA, USA) 90 min before treatment. After 30 min, myotubes were washed with phosphate-buffered saline (PBS), loaded with 10 mM CM-H2DCFDA (Invitrogen), and incubated for 1 h. Subsequently, myotubes were washed with PBS and exposed to acetaldehyde or acrolein followed by 20 min of incubation at 37 1C. Then, the myotubes were examined with an Axiovert 135 fluorescence inverted microscope (Carl Zeiss, Oberkochen, Germany) and the fluorescence intensity of DCF was measured using ImageJ software (National Institutes of Health (NIH), Bethesda, MD, USA). Fluorescence intensity was expressed as a percentage of the values of control myotubes (corresponding to 100%).
192
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
Cell lysates and Western blot analysis
Measurement of myotube diameter
After treatments with acetaldehyde or acrolein, myotubes were lysed for cytosolic proteins as previously described [30]. Myotubes were washed twice with PBS and lysed using lysis buffer (50 mM Tris– HCl, pH 7.4, 300 mM NaCl, 1.5 mM MgCl2, 200 mM EDTA, 0.1% Triton X-100) and 40 diluted protease and phosphatase inhibitors (Sigma– Aldrich) and centrifuged at 14,000 rpm for 10 min. Supernatants containing cytosolic proteins were collected and kept at −80 1C. Cytosolic proteins (20 μg) were loaded in each lane and separated by SDS–PAGE. Next, the proteins were transferred to nitrocellulose membranes. Membranes were blocked with 5% nonfat milk powder in Tris-buffered saline–0.125% Tween (TBS-T; Sigma–Aldrich) for 1 h and exposed overnight at 4 1C to primary antibodies. Primary antibodies against the following proteins were used: MyHC (1:1000), MAFbx/atrogin-1 (1:1000), MuRF1 (1:1000) (Santa Cruz Biotechnology, Santa Cruz, CA, USA); actin (1:4000) (Millipore, Billerica, MA, USA); p38 MAPK (1:1000), phospho-p38 MAPK (1:1000) (R&D Systems, Minneapolis, MN, USA). The next day, membranes were washed with TBS-T followed by 1 h incubation at ambient temperature with appropriate secondary antibodies (Jackson ImmunoResearch, West Grove, PA, USA). Detection was performed by enzyme-linked chemiluminescence (ECL; Biological Industries) using an ImageQuant LAS 4000 digital imager system (GE Healthcare, Chalfont St. Giles, Buckinghamshire, UK). Protein quantities were determined by densitometry and analyzed using Total Lab Software V2006C (Nonlinear Dynamics, Newcastle-on-Tyne, UK).
To assess the effects of acetaldehyde and acrolein on cell atrophy, diameter of myotubes was measured after treatments. Myotubes were examined using a phase-contrast microscope (CK40-SLP; Olympus, Tokyo, Japan; objective 20) and digital camera (UC30 Olympus, Japan). Three fields of view from three independent experiments were chosen randomly and the 10 largest myotubes in each field were measured in a blinded fashion using ImageJ software (NIH). Results were expressed as a percentage of the values of control myotubes (corresponding to 100%).
Protein loading control by Ponceau S staining In this study, the effects of acrolein and acetaldehyde on the main contractile muscle proteins MyHC and actin were studied. Therefore, actin could not be used as a housekeeping protein for loading control. As an alternative, quantification of total proteins by reversible Ponceau staining was used for protein loading control as previously described [30]. Briefly, before antibody probing, membranes were rinsed in Ponceau S solution (Bio-Rad, Hercules, CA, USA) for 10 min, followed by a brief rinse in double-distilled water until bands were clearly visible. In each lane, total protein quantities were determined by densitometry and used for normalization of ECL-detected proteins. Quantification of Ponceau staining before antibody probing has been previously validated as an alternative to actin blotting [33]. Real-time PCR RNA purification from myotubes was performed using a High Pure RNA isolation kit (Roche, Mannheim, Germany) according to the manufacturer's instructions. RNA concentrations were quantified at 260 nm by a Nanodrop spectrophotometer (Nanodrop Technologies, Wilmington, DE, USA). Samples of 1 μg RNA were used to synthesize cDNA using a High Capacity cDNA reverse transcription kit (Applied Biosystems, Foster City, CA, USA). QPCR was performed as previously described [30] using a qPCR SYBR Green ROX mix (Thermo Scientific, Basingstoke, UK) and Corbett Rotor-Gene 6000 (Qiagen, Hilden, Germany). For each sample, a value of the threshold cycle (Ct) was calculated using Rotor Gene 6000 series software (Qiagen) based on the time changes in mRNA expression level calculated subsequent to normalization with glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The abundance of target mRNA relative to GAPDH was determined by the ΔΔCt relative quantification method. The following primers were used (Sigma–Aldrich): GAPDH forward, 5′-AGGTCGGTGTGAACGGATTTG-3′, reverse, 5′-TGTAGACCATGTAGTTGAGGTCA-3′; MAFbx/atrogin-1 forward, 5′-CAGCTTCGTGAGCGACCTC-3′, reverse, 5′-GCAGTCGAGAAGTCCAGTC-3′; and MuRF1 forward, 5′-GTGTGAGGTGCCTACTTGCTC-3′, reverse, 5′-GCTCAGTCTTCTGTCCTTGGA-3′.
Statistical analysis Results were expressed as means + standard error (SE) of three independent experiments. Student's t test and one-way ANOVA followed by Tukey's or Dunnett's tests were used for statistical analysis using SPSS 16 software (IBM, USA). A p o 0.05 was considered statistically significant.
Results The effects of acetaldehyde on viability of myotubes To examine the effects of acetaldehyde on catabolism of C2 myotubes, it was essential to assess its effects on viability of myotubes and to find working concentrations at which acetaldehyde would not be cytotoxic. Therefore, differentiated myotubes were exposed to increasing levels of acetaldehyde (up to 500 mM) and incubated at 37 1C. After 24 h, viability of the myotubes was assessed by MTT assay as described under Materials and methods. Exposure of C2 myotubes to acetaldehyde at all concentrations examined did not cause a significant reduction in viability of the myotubes. The effects of acetaldehyde on the viability of myotubes at concentrations of 100, 300, and 500 mM are presented in Fig. 1. The effects of acetaldehyde on ROS generation Recently we have shown that shortly after exposure to CS, generation of ROS in C2 myotubes was increased and later led to muscle catabolism [30]. To assess the role of acetaldehyde in muscle catabolism it was of interest to explore its effects on ROS generation in C2 myotubes. Therefore, differentiated myotubes were exposed to acetaldehyde at increasing levels (up to 500 mM) in the presence or absence of 2 mM NAC and incubated at 37 1C. Twenty minutes after incubation, ROS generation was assessed by
Fig. 1. The effects of acetaldehyde on viability of myotubes. Myotubes were exposed to increasing levels of acetaldehyde and incubated for 24 h. After incubation, viability of the myotubes was measured by MTT assay and compared with untreated myotubes. Results are expressed as means + SE of three different experiments.
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
193
Fig. 2. The effects of acetaldehyde on ROS generation. (A) Untreated myotubes. (B) Myotubes pretreated with vehicle and exposed to 500 mM acetaldehyde. (C) Myotubes pretreated with 2 mM NAC 1 h before exposure to 500 mM acetaldehyde. (D) Changes in fluorescence intensity representing ROS production are expressed as a percentage of the fluorescence intensity found in untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments.
DCF assay as described under Materials and methods. At all concentrations examined, exposure of C2 myotubes to acetaldehyde did not cause a significant increase in ROS production compared with untreated myotubes. Pretreatment with 2 mM NAC before acetaldehyde exposure did not affect ROS generation significantly. The effects of acetaldehyde on ROS generation at the maximal concentration of 500 mM in the presence or absence of NAC are presented in Fig. 2.
Atrogin-1 MuRF1 Acetaldehyde concentration (µM)
CTL
100
300
500
The effects of acetaldehyde on muscle-specific E3 ligases Muscle catabolism is known to be mediated by upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1. To investigate the effects of acetaldehyde on atrogin-1 and MuRF1, C2 myotubes were exposed to acetaldehyde at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After incubation with acetaldehyde, protein levels of atrogin-1 and MuRF1 were examined by Western blot as described under Materials and methods. At all concentrations and time points examined, no significant change in protein levels of atrogin-1 or MuRF1 was found after exposure to acetaldehyde. The effects of 24 h exposure to acetaldehyde on protein levels of atrogin-1 and MuRF1 at concentrations of 100, 300, and 500 mM are presented in Fig. 3. The effects of acetaldehyde on muscle contractile proteins and diameter of myotubes To examine the effects of acetaldehyde on the main contractile muscle proteins MyHC and actin, C2 myotubes were exposed to acetaldehyde at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After incubation with acetaldehyde, protein levels of MyHC and actin were examined by Western blot. At all concentrations and time points examined, no significant change in protein levels of MyHC and actin was found after exposure to acetaldehyde. The effects of 24 h exposure
Fig. 3. The effects of acetaldehyde on muscle-specific E3 ligases. Myotubes were incubated with increasing concentrations of acetaldehyde (up to 500 mM) for 24 h. Untreated myotubes served as control. After incubation, cell lysates were prepared and subjected to Western blot analysis with antibodies against atrogin-1 and MuRF1. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are expressed as means + SE of three independent experiments.
to acetaldehyde on protein levels of MyHC and actin at concentrations of 100, 300, and 500 mM are presented in Fig. 4A. In addition, after incubation with acetaldehyde, myotubes were photographed and diameters were measured and compared with untreated myotubes as described under Materials and methods. Similarly, at all concentrations and time points examined, no significant change in the diameter of myotubes was found after exposure to acetaldehyde. Representative images of myotubes exposed to the
194
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
MyHC Actin Acetaldehyde concentration (µM)
CTL
100
300
500
Fig. 4. The effects of acetaldehyde on muscle contractile proteins and diameter of myotubes. (A) Myotubes were incubated with increasing concentrations of acetaldehyde (up to 500 mM) for 24 h. Untreated myotubes served as control. After incubation, cell lysates were prepared and subjected to Western blot analysis with antibodies against MyHC and actin. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are expressed as means + SE of three independent experiments. (B) Untreated myotubes. (C) Myotubes exposed to 500 mM acetaldehyde for 24 h. (D) Changes in diameter of myotubes are expressed as percentage of the diameter of untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments.
maximal concentration of 500 mM for 24 h in comparison with untreated myotubes are presented in Fig. 4B–D.
The effects of acrolein on viability of myotubes Acrolein is known as an extremely toxic unsaturated aldehyde [7]. Therefore, examining the effects of acrolein on the viability of C2 myotubes was essential to find a suitable concentration for experiments of acrolein exposure. Thus, myotubes were exposed to acrolein at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After exposure, the viability
of myotubes was assessed by MTT assay as described under Materials and methods. After 1 h incubation, exposure to acrolein at concentrations up 100 mM did not cause a significant reduction in cell viability. However, exposure to acrolein at 250 and 500 mM caused a significant reduction in the viability of myotubes (Fig. 5A). Subsequently, the effects of 25 mM acrolein on the viability of myotubes at increasing incubation periods were examined. After 6 h of incubation with 25 mM acrolein, the viability of myotubes decreased significantly. However, at 6 h, the viability of myotubes remained higher than 80% (Fig. 5B). Therefore, experiments were carried out with acrolein at a concentration of 25 mM for up to 6 h.
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
195
Acrolein increases ROS generation in myotubes Myotubes were exposed to 25 mM acrolein in the presence or absence of 2 mM NAC and incubated at 37 1C. After 20 min, ROS generation was assessed by the DCF assay. Exposure of C2 myotubes to acrolein caused a significant increase in ROS generation in comparison with untreated myotubes. Pretreatment with 2 mM NAC 1 h before acrolein exposure prevented acrolein-induced increase in ROS generation. Representative images of myotubes exposed to 25 mM acrolein in the presence or absence of NAC and compared with untreated myotubes are presented in Fig. 6. Acrolein upregulates muscle-specific E3 ligases To investigate the effects of acrolein on the expression of atrogin-1 and MuRF1, C2 myotubes were exposed to 25 mM acrolein and incubated for increasing time periods (up to 6 h). After incubation, protein levels of atrogin-1 and MuRF1 were examined by Western blot and mRNA levels were determined by qPCR as described under Materials and methods. A significant increase in mRNA levels of atrogin-1 and MuRF1 compared with untreated myotubes was detected at 1 and 2 h of incubation with acrolein (Fig. 7A). In addition, a significant increase in protein levels of atrogin-1 and MuRF1 was detected from 1 h until 6 h of incubation with acrolein (Fig. 7B). Acrolein induces MyHC degradation and reduction in diameter of myotubes
Fig. 5. The effects of acrolein on viability of myotubes. (A) Myotubes were exposed to increasing levels of acrolein and incubated for 1 h. (B) Myotubes were exposed to acrolein at 25 mM for increasing incubation periods. After incubation, the viability of myotubes was measured by MTT assay and compared with that of untreated myotubes. Results are expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
To examine the effects of acrolein on the atrophy of myotubes and the main contractile muscle proteins MyHC and actin, myotubes were exposed to 25 mM acrolein and incubated for increasing time intervals (up to 6 h). After incubation, myotubes were photographed and diameters were measured and compared with those of untreated myotubes. Subsequently, myotubes were lysed
Fig. 6. Acrolein increases ROS generation in myotubes. (A) Untreated myotubes. (B) Myotubes pretreated with vehicle and exposed to 25 mM acrolein. (C) Myotubes pretreated with 2 mM NAC 1 h before exposure to 25 mM acrolein. (D) Changes in fluorescence intensity representing ROS production are expressed as percentage of the fluorescence intensity found in untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
196
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
myotubes were pretreated with 5 μM p38 MAPK inhibitor SB203580 or vehicle 15 min before exposure to 25 mM acrolein and incubation (up to 6 h). After incubation, phosphorylation of p38 MAPK and protein levels of atrogin-1, MuRF1, MyHC, and actin were examined by Western blot. Significant phosphorylation of p38 MAPK was observed at 1 and 2 h of exposure to acrolein. Pretreatment with SB203580 blocked acrolein-induced activation of p38 MAPK (Fig. 9A). In addition, inhibition of p38 MAPK abolished acrolein-induced increase in protein levels of atrogin-1 and MuRF1 and degradation of MyHC at 2 h of exposure to acrolein. Representative blots are presented in Fig. 9B and C.
Discussion
Atrogin -1 MuRF1 Acrolein (25 µM) incubation time
C T L 1 hr
2 hr
6 hr
Fig. 7. Acrolein upregulates muscle-specific E3 ligases. Myotubes were exposed to 25 mM acrolein and incubated for increasing time periods. Untreated myotubes served as control. (A) After incubation, RNA was isolated and subjected to reverse transcription and qPCR analysis to determine the expression of atrogin-1 and MuRF1. Data were normalized to GAPDH expression and are relative to the corresponding value of control myotubes. (B) After incubation with acrolein, cell lysates were prepared and subjected to Western blot analysis with antibodies against atrogin-1 and MuRF1. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control myotubes. Results are expressed as means + SE of three independent experiments. *p o 0.05 versus control myotubes.
and subjected to Western blot analysis. Diameter of myotubes decreased depending on acrolein incubation time. A significant decrease in diameter was found at 2 and 6 h of incubation with acrolein compared with untreated myotubes. Representative images are presented in Fig. 8A–E. Also, protein levels of MyHC decreased depending on acrolein incubation times. A significant decrease in MyHC was found at 2 and 6 h of incubation with acrolein compared with untreated myotubes. The protein level of actin did not change significantly. Representative blots are presented in Fig. 8F. The involvement of p38 MAPK in acrolein-induced muscle catabolism In previous studies, activation of p38 MAPK was shown to be one of the initial steps for CS-induced catabolism of skeletal muscle [14,30,34]. Moreover, inhibition of p38 MAPK abolished the catabolic effects caused by exposure to CS [30] or extract of CS [34]. Therefore, it was of interest to examine the role of p38 MAPK in acrolein-induced catabolism of myotubes. Thus,
In this study, skeletal myotubes were exposed to various levels of acetaldehyde and acrolein to assess their role in catabolism of skeletal muscle. For the first time, it was shown that exposure of skeletal myotubes to acrolein and not acetaldehyde resulted in cell atrophy and breakdown of MyHC through increased oxidative stress, activation of p38 MAPK, and upregulation of the musclespecific E3 ligases atrogin-1 and MuRF1. Several conditions in which levels of acetaldehyde and acrolein may be increased were associated with muscle catabolism. Acetaldehyde was previously suggested to damage skeletal muscle in alcoholic myopathy [3,4]. In addition, acetaldehyde is one of the major constituents of CS [19]. Epidemiological, clinical, in vivo, and in vitro studies reported that smoking or exposure to CS can damage skeletal muscle and promote its catabolism [14,25–31,34]. Therefore, it was of great interest to study the effects of acetaldehyde exposure on muscle catabolism in skeletal myotubes. Blood levels of acetaldehyde in healthy subjects were reported at 12.2 mM [35]. After administration of alcohol, mean concentrations of acetaldehyde in alcoholics were 42.7 mM in comparison with 26.5 mM in nonalcoholics [36]. Smoking a single cigarette can elevate blood levels of acetaldehyde by up to 9.5 mM [19]. In this study, the catabolic effects of acetaldehyde were investigated in physiological and supraphysiological levels of up to 500 mM. At all concentrations and time points examined, acetaldehyde was not cytotoxic and did not promote catabolism of skeletal myotubes. C2 myotubes exposed to acetaldehyde in concentrations of up to 500 mM did not demonstrate increased ROS generation or upregulation of atrogin-1 and MuRF1. Also, no significant reductions in the diameter of myotubes or in the levels of the main contractile muscle proteins MyHC and actin were found. It was previously shown that exposure of human myocytes to 200 mM acetaldehyde decreased basal protein synthesis without altering the rate of protein degradation [5]. Also, injection of acetaldehyde into rats resulted in reduced fractional rates of protein synthesis in skeletal muscles [6]. Therefore, it is possible that acetaldehyde may promote catabolism of skeletal muscle, not through the activation of catabolic pathways but through inhibition of anabolic pathways. To prove this hypothesis, future studies should focus on the effects of acetaldehyde on inhibition of anabolic pathways in skeletal muscle. These pathways include the insulin-like growth factor 1/phosphatidylinositol 3-kinase/protein kinase B and the mammalian target of rapamycin signaling pathways, which regulate protein synthesis in skeletal muscle [37]. Acrolein has numerous biological effects including regulation of signaling molecules and activation of transcription factors [7]. Conditions in which blood or tissue levels of acrolein may be increased include renal failure, oxidative stress, and heavy smoking [10,11]. During inflammation and oxidative stress acrolein can be produced endogenously via myeloperoxidase-mediated degradation of threonine and amine oxidase-mediated degradation of spermine and spermidine [9]. Wasting of skeletal muscle is
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
197
MyHC Actin Acrolein (25 µM) incubation time
CTL
1 hr
2 hr
6 hr
Fig. 8. Acrolein induces MyHC degradation and reduction in diameter of myotubes. (A) Untreated myotubes. (B) Myotubes exposed to 25 mM acrolein for 1 h. (C) Myotubes exposed to 25 mM acrolein for 2 h. (D) Myotubes exposed to 25 mM acrolein for 6 h. (E) Changes in diameter of myotubes are expressed as a percentage of the diameter of untreated myotubes. (F) Cell lysates were prepared and subjected to Western blot analysis with antibodies against MyHC and actin. Protein levels were normalized by total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are relative to control and expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
common in renal failure patients [13] and may be increased in situations of oxidative stress and chronic inflammation [14]. The main source of exogenous acrolein in humans is exposure to CS [2,9]. CS is considered to make up a large proportion of total human exposure to acrolein [9]. As mentioned earlier, the catabolic effects of CS were demonstrated previously [14,25–31,34]. Therefore, it was of great interest to assess the effects of acrolein exposure on catabolism of skeletal myotubes. To the best of our knowledge, the effects of acrolein exposure on skeletal muscle have not been studied previously. However, the effects of acrolein on cardiac muscle were investigated earlier. Ismahil et al. [15] found that chronic oral exposure of mice to acrolein (1 mg/kg) resulted in cardiomyopathy accompanied by contractile dysfunction and myocardial oxidative stress. Also, Luo et al. [1] showed that intravenous injection of acrolein (0.5 mg/kg) into mice resulted in myocardial dysfunction and increased acrolein–protein adducts, including adducts to sarcomeric proteins such as myosin light polypeptide. The above studies demonstrate that acrolein administered both orally and intravenously can reach cardiac muscle tissue and promote cardiomyopathy. Previous studies investigated the effects of acrolein on smooth muscle cultures and demonstrated its role in activation of signaling pathways and gene regulation. Ranganna et al. [16] found that exposure of rat vascular smooth muscle cells to acrolein resulted in activation of several members of the MAPK family, including p38 MAPK. A concentration- and time-dependent activation of p38 MAPK was induced by acrolein and was inhibited by SB203580 pretreatment. Volpi et al. [17] exposed human airway smooth muscle cells to CS extract or acrolein at concentrations of 10–100 mM. They found that acrolein mimicked the effects of CS
extract and caused activation of p38 MAPK, which led to upregulation of vascular endothelial growth factor. Acrolein at 60 mM elicited a rapid phosphorylation of p38 MAPK that peaked at 1 h and was blocked by SB203580. Similarly, Moretto et al. [18] exposed human bronchial smooth muscle cells to CS extract or acrolein at concentrations of 10–60 mM. Both CS extract and acrolein induced p38 MAPK phosphorylation, which resulted in upregulation of interleukin-8. Peak phosphorylation was found at 30 min of exposure to 30 mM acrolein and was abolished by SB203580. In this study, it is shown that exposure of C2 myotubes to acrolein at the level of 25 mM caused atrophy of myotubes and breakdown of MyHC via increased ROS generation, activation of p38 MAPK, and upregulation of atrogin-1 and MuRF1. Similar to the above studies, peak phosphorylation of p38 MAPK was observed at 1 h of exposure to acrolein and was prevented by SB203580 pretreatment. Recently, we have shown that CS exposure to skeletal myotubes also caused cell atrophy and degradation of MyHC through increased oxidative stress, activation of p38 MAPK, and upregulation of atrogin-1 and MuRF1. Inhibition of p38 MAPK prevented CS-induced muscle catabolism in skeletal myotubes [30]. In this study, acrolein was shown to mimic the effects of CS by activating a similar signaling cascade that resulted in muscle catabolism. Because acrolein mimicked the effects of CS on skeletal muscle, it may be suggested that acrolein is one of the main components of CS responsible for its catabolic effects. However, CS is a complex aerosol consisting of thousands of constituents [38]. Various constituents of CS have the potential to induce catabolism of skeletal muscle [31]. For instance, CS is a major source of both ROS
198
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
P-p38 p38 Acrolein (25 µM) CTL incubation time
1 hr
1 hr
2 hr
2 hr
6 hr
6 hr
SB203580
-
+
-
+
-
+
-
Atrogin-1
MyHC
MuRF1
Actin
Acrolein (25 µM) incubation time SB203580
CTL
2 hr
2 hr
-
-
+
Acrolein (25 µM) incubation time SB203580
CTL
2 hr
2 hr
-
-
+
Fig. 9. The involvement of p38 MAPK in acrolein-induced muscle catabolism. Myotubes were exposed to 25 mM acrolein in the presence of 5 μM SB203580 or vehicle. Untreated myotubes served as control. After incubation with acrolein, cell lysates were prepared and subjected to Western blot analysis with antibodies against p38 MAPK (p38), phosphorylated p38 MAPK (P-p38), atrogin-1, MuRF1, MyHC, and actin. (A) Protein levels of P-p38 and p38 were quantified by densitometry and values of P-p38 were normalized to p38 and compared with control. (B and C) Protein levels were normalized by total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control myotubes. Results are expressed as means + SE of three independent experiments. *p o 0.05 versus control myotubes.
and reactive nitrogen species (RNS) [31]. ROS present in CS, such as hydrogen peroxide (H2O2), were shown to upregulate atrogin-1 and MuRF1 and stimulate ubiquitination of muscle proteins in skeletal myotubes [39]. Also, RNS present in CS, such as peroxynitrite (ONOO−), were demonstrated to cause degradation of muscle proteins in cultured myotubes mediated by activation of nuclear factor-κB [40]. Therefore, it seems that in addition to acrolein, ROS and RNS from CS may also play an important role in the catabolism of skeletal muscle in smokers. Our findings demonstrate that acrolein-induced catabolism of skeletal muscle is related to increased oxidative stress as found by the DCF assay (Fig. 6). Acrolein may increase ROS production and oxidative stress in several mechanisms. First, acrolein can activate the nicotinamide adenine dinucleotide phosphate oxidase complex, leading to superoxide production. Superoxide dismutase converts superoxide into H2O2 and activates inducible nitric oxide synthase, leading to production of nitric oxide and formation of ONOO−, which ultimately causes oxidative and nitrosative stress [41]. In addition, acrolein can react with cellular nucleophiles, such as reduced glutathione (GSH) [41]. The conjugation of acrolein with GSH depletes its cellular levels, impairing the clearance of free radicals that results in increased oxidative stress [16,41].
The mechanisms by which acrolein increases oxidative stress in skeletal muscle should be investigated in future studies. This study has certain limitations that need to be addressed when comparing in vitro exposure of acrolein with clinical conditions of increased exposure to acrolein. First, myotubes were treated with acrolein prepared in low-protein medium consisting of 2% horse serum as required for their successful differentiation from myoblasts into myotubes. However, human plasma contains a higher percentage of protein (approximately 6–8%). Thus, a higher percentage of protein in the medium as in plasma might have a different impact on the cytotoxicity and biological effects of acrolein that were found in this study. In addition, the working concentration of acrolein (25 mM) was determined from viability assays that examined the effects of acrolein on the viability of myotubes at increasing concentrations of up to 500 mM. Another limitation is the difficulty in comparing the concentrations of acrolein used in our in vitro study with its physiological levels in various conditions. For instance, during inflammation, phagocytes can generate acrolein by the action of myeloperoxidase on threonine that can circulate in high concentrations of about 2 mM [10]. In renal failure patients, plasma levels of acrolein were studied previously. Using high-performance
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
liquid chromatography (HPLC), Igarashi et al. [12] found that free acrolein in plasma of uremic patients was 1.4 mM, in comparison with 0.5 mM in control subjects. Acrolein found as protein conjugates in uremic patients was equivalent to 170 mM, about fivefold higher than its concentration in the plasma of control subjects. Thus, in comparison with free acrolein levels found in plasma of renal failure patients, the acrolein concentration of 25 mM used in this study appears to be supraphysiological. In heavy smokers, circulating or tissue concentrations of acrolein are increased [10,11]. However, as far as we are aware, the exact concentrations of acrolein in the plasma of smokers and the levels to which skeletal muscles of smokers are exposed to acrolein have not been studied yet. Annovazzi et al. [23] performed HPLC analysis on the saliva of smokers versus nonsmokers and showed that average acrolein concentrations in heavy smokers were 21 mM compared with 2.3 mM in nonsmokers. Eiserich et al. [21] calculated that the concentration of acrolein in respiratory tract lining fluids (RTLF) after exposure to one cigarette is 80 mM. Carmella et al. [24] found that the levels of 3,3-hydroxypropylmercapturic acid, a major acrolein metabolite formed by GSH conjugation, in urine are related to cigarette smoking, suggesting that acrolein is transported from the lung into the systemic circulation. The calculation by Eiserich et al. [21] suggests that the acrolein concentration of 25 mM used in this study may be within the physiological range. However, this calculation was based on the assumption that all of the components of CS are completely deposited within the RTLF with no further metabolism. Thus, RTLF and plasma concentrations of acrolein after exposure to one cigarette may be lower than calculated. On the other hand, the above calculation was based on experiments that examined exposure to a single cigarette without considering the effects of prolonged exposure to acrolein as in heavy smokers. Therefore, until the exact level of acrolein to which skeletal muscles of smokers are exposed is investigated, it will be difficult to compare the concentrations of acrolein used in our in vitro study with its physiological levels in smokers.
Conclusions Our findings demonstrate that exposure of skeletal myotubes to acrolein and not acetaldehyde resulted in atrophy of myotubes and breakdown of MyHC via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1. Although within the limitations of an in vitro study, these findings may provide a molecular explanation for muscle wasting in conditions of increased exposure to acrolein, including cigarette smoking, renal failure, oxidative stress, and inflammation. Future clinical studies should examine the role of acrolein in wasting and dysfunction of skeletal muscle in these pathologies and in heavy smokers.
Acknowledgments This study was supported by grants from the Rappaport Institute, the Krol Foundation of Barnegat, New Jersey, the Myers–JDC Brookdale Institute of Gerontology and Human Development, and ESHEL—the association for planning and development of services for the aged in Israel. Special thanks to Dr. Alvira Bromosov for her help in the DCF assay.
199
References [1] Luo, J.; Hill, B. G.; Gu, Y.; Cai, J.; Srivastava, S.; Bhatnagar, A.; Prabhu, S. D. Mechanisms of acrolein-induced myocardial dysfunction: implications for environmental and endogenous aldehyde exposure. Am. J. Physiol. Heart Circ. Physiol 293:H3673–H3684; 2007. [2] Voulgaridou, G. P.; Anestopoulos, I.; Franco, R.; Panayiotidis, M. I.; Pappa, A. DNA damage induced by endogenous aldehydes: current state of knowledge. Mutat. Res. 711:13–27; 2011. [3] Preedy, V. R.; Keating, J. W.; Peters, T. J. The acute effects of ethanol and acetaldehyde on rates of protein synthesis in type I and type II fiber-rich skeletal muscles of the rat. Alcohol Alcohol 27:241–251; 1992. [4] Preedy, V. R.; Adachi, J.; Ueno, Y.; Ahmed, S.; Mantle, D.; Mullatti, N.; Rajendram, R.; Peters, T. J. Alcoholic skeletal muscle myopathy: definitions, features, contribution of neuropathy, impact and diagnosis. Eur. J. Neurol. 8:677–687; 2001. [5] Hong-Brown, L. Q.; Frost, R. A.; Lang, C. H. Alcohol impairs protein synthesis and degradation in cultured skeletal muscle cells. Alcohol Clin. Exp. Res. 25:1373–1382; 2001. [6] Oba, T.; Maeno, Y.; Ishida, K. Differential contribution of clinical amounts of acetaldehyde to skeletal and cardiac muscle dysfunction in alcoholic myopathy. Curr. Pharm. Des 11; 2005. 791-780. [7] Berry, K. A.; Henson, P. M.; Murphy, R. C. Effects of acrolein on leukotriene biosynthesis in human neutrophils. Chem. Res. Toxicol. 21:2424–2432; 2008. [8] Anderson, E. J.; Katunga, L. A.; Willis, M. S. Mitochondria as a source and target of lipid peroxidation products in healthy and diseased heart. Clin. Exp. Pharmacol. Physiol. 39:179–193; 2012. [9] Stevens, J. F.; Maier, C. S. Acrolein: sources, metabolism, and biomolecular interactions relevant to human health and disease. Mol. Nutr. Food Res. 52:7–25; 2008. [10] Gugliucci, A. Antithrombin activity is inhibited by acrolein and homocysteine thiolactone: protection by cysteine. Life Sci 82:413–418; 2008. [11] Misonou, Y.; Asahi, M.; Yokoe, S.; Miyoshi, E.; Taniguchi, N. Acrolein produces nitric oxide through the elevation of intracellular calcium levels to induce apoptosis in human umbilical vein endothelial cells: implications for smoke angiopathy. Nitric Oxide 14:180–187; 2006. [12] Igarashi, K.; Ueda, S.; Yoshida, K.; Kashiwagi, K. Polyamines in renal failure. Amino Acids 31:477–483; 2006. [13] Sukhanov, S.; Semprun-Prieto, L.; Yoshida, T.; Tabony, A. M.; Higashi, Y.; Galvez, S.; Delafontaine, P. Angiotensin II, oxidative stress and skeletal muscle wasting. Am. J. Med. Sci. 342:143–147; 2011. [14] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Sarcopenia and smoking: a possible cellular model of cigarette smoke effects on muscle protein breakdown. Ann. N. Y. Acad. Sci 1259:47–53; 2011. [15] Ismahil, M. A.; Hamid, T.; Haberzettl, P.; Gu, Y.; Chandrasekar, B.; Srivastava, S.; Bhatnagar, A.; Prabhu, S. D. Chronic oral exposure to the aldehyde pollutant acrolein induces dilated cardiomyopathy. Am. J. Physiol. Heart Circ. Physiol 301:2050–2060; 2011. [16] Ranganna, K.; Yousefipour, Z.; Nasif, R.; Yatsu, F. M.; Milton, S. G.; Hayes, B. E. Acrolein activates mitogen-activated protein kinase signal transduction pathways in rat vascular smooth muscle cells. Mol. Cell. Biochem. 240:83–98; 2002. [17] Volpi, G.; Facchinetti, F.; Moretto, N.; Civelli, M.; Patacchini, R. Cigarette smoke and α,β-unsaturated aldehydes elicit VEGF release through the p38 MAPK pathway in human airway smooth muscle cells and lung fibroblasts. Br. J. Pharmacol 163:649–661; 2011. [18] Moretto, N.; Facchinetti, F.; Southworth, T.; Civelli, M.; Singh, D.; Patacchini, R. α,β-Unsaturated aldehydes contained in cigarette smoke elicit IL-8 release in pulmonary cells through mitogen-activated protein kinases. Am. J. Physiol. Lung Cell. Mol. Physiol 296:839–848; 2009. [19] Talhout, R.; Opperhuizen, A.; Van Amsterdam, J. G. Role of acetaldehyde in tobacco smoke addiction. Eur. Neuropsychopharmacol. 17:627–636; 2007. [20] Smith, C. J.; Fischer, T. H. Particulate and vapor phase constituents of cigarette mainstream smoke and risk of myocardial infarction. Atherosclerosis 158: 257–267; 2011. [21] Eiserich, J. P.; Van der Vliet, A.; Handelman, G. J.; Halliwell, B.; Cross, C. E. Dietary antioxidants and cigarette smoke-induced biomolecular damage: a complex interaction. Am. J. Clin. Nutr. 62:1490S–1500S; 1995. [22] Colombo, G.; Aldini, G.; Orioli, M.; Giustarini, D.; Gornati, R.; Rossi, R.; Colombo, R.; Carini, M.; Milzani, A.; Dalle-Donne, I. Water-soluble alpha, beta-unsaturated aldehydes of cigarette smoke induce carbonylation of human serum albumin. Antioxid. Redox Signaling 12:349–364; 2010. [23] Annovazzi, L.; Cattaneo, V.; Viglio, S.; Perani, E.; Zanone, C.; Rota, C.; Pecora, F.; Cetta, G.; Silvestri, M.; Iadarola, P. High-performance liquid chromatography and capillary electrophoresis: methodological challenges for the determination of biologically relevant low-aliphatic aldehydes in human saliva. Electrophoresis 25:1255–1263; 2004. [24] Carmella, S. G.; Chen, M.; Zhang, Y.; Zhang, S.; Hatsukami, D. K.; Hecht, S. S. Quantitation of acrolein-derived (3-hydroxypropyl)mercapturic acid in human urine by liquid chromatography–atmospheric pressure chemical ionization tandem mass spectrometry: effects of cigarette smoking. Chem. Res. Toxicol. 20:986–990; 2007. [25] Petersen, A. M.; Magkos, F.; Atherton, P.; Selby, A.; Smith, K.; Rennie, M. J.; Pedersen, B. K.; Mittendorfer, B. Smoking impairs muscle protein synthesis and increases the expression of myostatin and MAFbx in muscle. Am. J. Physiol. Endocrinol. Metab. 293:843–848; 2007.
200
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
[26] Montes de Oca, M.; Loeb, E.; Torres, S. H.; De Sanctis, J.; Hernández, N.; Tálamo, C. Peripheral muscle alterations in non-COPD smokers. Chest 133:13–18; 2008. [27] Castillo, E. M.; Goodman-Gruen, D.; Kritz-Silverstein, D.; Morton, D. J.; Wingard, D. L.; Barrett-Connor, E. Sarcopenia in elderly men and women: the Rancho Bernardo study. Am. J. Prev. Med. 25:226–231; 2003. [28] Szulc, P.; Duboeuf, F.; Marchand, F.; Delmas, P. D. Hormonal and lifestyle determinants of appendicular skeletal muscle mass in men: the MINOS study. Am. J. Clin. Nutr. 80:496–503; 2004. [29] Lee, J. S.; Auyeung, T. W.; Kwok, T.; Lau, E. M.; Leung, P. C.; Woo, J. Associated factors and health impact of sarcopenia in older Chinese men and women: a cross-sectional study. Gerontology 53:404–410; 2007. [30] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Cigarette smoke and muscle catabolism in C2 myotubes. Mech. Ageing Dev. 134; 2013. 24-23. [31] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Identification of possible cigarette smoke constituents responsible for muscle catabolism. J. Muscle Res. Cell Motil. 33:199–208; 2012. [32] Sadowska, A. M.; Manuel-Y-Keenoy, B.; De Backer, W. A. Antioxidant and antiinflammatory efficacy of NAC in the treatment of COPD: discordant in vitro and in vivo dose-effects: a review. Pulm. Pharmacol. Ther 20:9–22; 2007. [33] Romero-Calvo, I.; Ocon, B.; Martinez-Moya, P.; Suarez, M. D.; Zarzuelo, A.; Martinez-Augustin, O.; Medina, F. S. Reversible Ponceau staining as a loading control alternative to actin in Western blots. Anal. Biochem. 401:318–320; 2010.
[34] Liu, Q.; Xu, W. G.; Luo, Y.; Han, F. F.; Yao, X. H.; Yang, T. Y.; Zhang, Y.; Pi, W. F.; Guo, X. J. Cigarette smoke-induced skeletal muscle atrophy is associated with up-regulation of USP-19 via p38 and ERK MAPKs. J. Cell. Biochem. 112:2307–2316; 2011. [35] Pezzoli, C.; Galli-kienle, M.; Padova, C.; Stramentinoli, G. HPLC method for the evaluation of blood acetaldehyde without ethanol interference. J. Liq. Chromatogr 7:765–778; 1984. [36] Korsten, M. A.; Matsuzaki, S.; Feinman, L.; Lieber, C. S. High blood acetaldehyde levels after ethanol administration: difference between alcoholic and nonalcoholic subjects. N. Engl. J. Med. 292:386–389; 1975. [37] Meng, S. J.; Yu, L. J. Oxidative stress, molecular inflammation and sarcopenia. Int. J. Mol. Sci. 11:1509–1526; 2010. [38] Swan, E. G.; Lessov-Schlaggar, C. N. The effects of tobacco smoke and nicotine on cognition and the brain. Neuropsychol. Rev. 17:259–273; 2007. [39] Li, Y.; Chen, Y.; Li, A. S.; Reid, M. B. Hydrogen peroxide stimulates ubiquitinconjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes. Am. J. Physiol. Cell Physiol 285:C806–C812; 2003. [40] Bar-Shai, M.; Reznick, A. Z. Reactive nitrogen species induce nuclear factor-κBmediated protein degradation in skeletal muscle cells. Free Radic. Biol. Med 40:2112–2125; 2006. [41] Ansari, M. A.; Keller, J. N.; Scheff, S. W. Protective effect of Pycnogenol in human neuroblastoma SH-SY5Y cells following acrolein-induced cytotoxicity. Free Radic. Biol. Med. 45:1510–1519; 2008.
Contents lists available at SciVerse ScienceDirect
Free Radical Biology and Medicine journal homepage: www.elsevier.com/locate/freeradbiomed
Original Contribution
The effects of acetaldehyde and acrolein on muscle catabolism in C2 myotubes Oren Rom a, Sharon Kaisari a, Dror Aizenbud a,b, Abraham Z. Reznick a,n a b
Department of Anatomy and Cell Biology, Rappaport Faculty of Medicine, Technion–Israel Institute of Technology, Bat Galim, Haifa 31096, Israel Orthodontic and Craniofacial Department, Rambam Health Care Campus, Bat Galim, Haifa 31096, Israel
art ic l e i nf o
a b s t r a c t
Article history: Received 24 February 2013 Received in revised form 23 May 2013 Accepted 12 June 2013 Available online 20 June 2013
The toxic aldehydes acetaldehyde and acrolein were previously suggested to damage skeletal muscle. Several conditions in which exposure to acetaldehyde and acrolein is increased were associated with muscle wasting and dysfunction. These include alcoholic myopathy, renal failure, oxidative stress, and inflammation. A main exogenous source of both acetaldehyde and acrolein is cigarette smoking, which was previously associated with increased muscle catabolism. Recently, we have shown that exposure of skeletal myotubes to cigarette smoke stimulated muscle catabolism via increased oxidative stress, activation of p38 MAPK, and upregulation of muscle-specific E3 ubiquitin ligases. In this study, we aimed to investigate the effects of acetaldehyde and acrolein on catabolism of skeletal muscle. Skeletal myotubes differentiated from the C2 myoblast cell line were exposed to acetaldehyde or acrolein and their effects on signaling pathways related to muscle catabolism were studied. Exposure of myotubes to acetaldehyde did not promote muscle catabolism. However, exposure to acrolein caused increased generation of free radicals, activation of p38 MAPK, upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1, degradation of myosin heavy chain, and atrophy of myotubes. Inhibition of p38 MAPK by SB203580 abolished acrolein-induced muscle catabolism. Our findings demonstrate that acrolein but not acetaldehyde activates a signaling cascade resulting in muscle catabolism in skeletal myotubes. Although within the limitations of an in vitro study, these findings indicate that acrolein may promote muscle wasting in conditions of increased exposure to this aldehyde. & 2013 Elsevier Inc. All rights reserved.
Keywords: Acetaldehyde Acrolein Muscle catabolism Oxidative stress p38 MAPK E3 ubiquitin ligases Free radicals
Aldehydes are highly reactive and toxic molecules. Exogenous exposure to aldehydes has been associated with increased risk of various pathologies and death [1]. Aldehydes are also produced endogenously during metabolism of amino acids and lipids and are involved in numerous biochemical and physiological processes. Acetaldehyde and acrolein are toxic aldehydes and their involvement in several pathologies has been studied extensively in the past [2]. The saturated aldehyde acetaldehyde is mainly produced by metabolism of alcohol in the liver and is considered to be responsible for the pathological complications of alcohol consumption [2]. Acetaldehyde has been previously suggested to damage skeletal muscle in alcoholism [3,4]. Approximately 50% of alcohol misusers suffer from chronic alcoholic myopathy in
Abbreviations: CS, cigarette smoke; MyHC, myosin heavy chain; MAPK, mitogenactivated protein kinase; atrogin-1, muscle atrophy F-box protein; MuRF1, muscle ring finger-1 protein; NAC, N-acetylcysteine; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; ROS, reactive oxygen species; RNS, reactive nitrogen species; GSH, reduced glutathione n Corresponding author. Fax: +97248295403. E-mail address:[email protected] (A.Z. Reznick). 0891-5849/$ - see front matter & 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.freeradbiomed.2013.06.024
which muscle mass may be reduced by up to 30%. In alcoholics, blood levels of acetaldehyde are relatively high and acetaldehyde was suggested to be a primary factor in alcohol-induced muscle damage [4]. Previous studies showed that both in vitro exposure of myocytes to acetaldehyde and injection of acetaldehyde to rats resulted in low synthesis of muscle proteins [5,6]. Acrolein is an extremely toxic and highly reactive α,β-unsaturated aldehyde that has been linked with various diseases and shown to regulate activation of transcription factors and signaling molecules [7]. Endogenous sources of acrolein include lipid peroxidation, in which highly reactive aldehydes are produced [2,8]. These aldehydes have been recognized as causative factors in aging and age-associated diseases, including cardiovascular disease and diabetes [8]. Acrolein can also be produced in situations of oxidative stress and inflammation during degradation of threonine by myeloperoxidase and degradation of spermine and spermidine by amine oxidase [9]. High serum or tissue levels of acrolein have been described in various pathological conditions such as renal failure, diabetes, and oxidative stress [10–12]. Wasting and dysfunction of skeletal muscle are common in both renal failure and diabetic patients [13]. Oxidative stress and chronic inflammation in which acrolein levels may be increased
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
are known to activate signaling pathways that lead to atrophy and loss of skeletal muscle fibers [9,14]. Thus, high levels of acrolein in these pathological conditions may exacerbate the catabolism of skeletal muscle. Previous studies have investigated the effects of acrolein exposure on cardiac and smooth muscle. In rodents, acrolein exposure was shown to induce cardiomyopathy characterized by increased oxidative stress and upregulation of proinflammatory cytokines [1,15]. In smooth muscle cultures treated with acrolein, activation of various signaling pathways, including the mitogen-activated protein kinase (MAPK) pathway, was demonstrated [16–18]. However, as far as we are aware, the effects of acrolein on skeletal muscle have not been examined previously. Cigarette smoke (CS) is a major exogenous source of both acetaldehyde and acrolein [2,7,9]. CS from a common cigarette contains approximately 1.5 mg of acetaldehyde. Non-proteinbound acetaldehyde can cross the alveolar–capillary membranes of the lungs and cigarette smoking can increase blood concentrations of acetaldehyde [19]. CS contains 60–100 mg of acrolein per cigarette [20]. Saliva, blood, and urine levels of acrolein and its metabolites are increased in smokers in comparison with nonsmokers [10,21–24]. Previous studies have revealed an association between smoking and skeletal muscle damage. Increased muscular damage and expression of genes associated with muscle catabolism were found in skeletal muscle biopsies taken from smokers and compared with nonsmokers [25,26]. In addition, smoking has been previously associated with sarcopenia, the age-related loss of muscle mass and strength [27–29]. Recently, we have shown that CS exposure to skeletal myotubes caused cell atrophy and degradation of myosin heavy chain (MyHC) via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ligases of the ubiquitin–proteasome system: muscle atrophy F-box protein (atrogin-1) and muscle ring finger-1 protein (MuRF1) [30]. Acetaldehyde and acrolein present in CS were previously suggested to possess potential catabolic effects on skeletal muscle [31]. Increased levels of acetaldehyde and acrolein are evident in various states associated with muscle catabolism, including oxidative stress, inflammation, renal failure [9–12], and alcoholism [2–4]. The aim of this study was to investigate the effects of acetaldehyde and acrolein on catabolism of skeletal myotubes. This was done by exposing C2 skeletal myotubes to various levels acetaldehyde and acrolein. We hypothesized that in a manner similar to that of CS, exposure of cultured myotubes to acetaldehyde and acrolein will lead to muscle atrophy and degradation of muscle proteins via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ubiquitin ligases atrogin-1 and MuRF1.
Materials and methods Cell culture The C2 cell line of mouse skeletal myoblasts was a generous gift from Professor Eyal Bengal (Rappaport Faculty of Medicine, Technion, Israel). C2 myoblasts were grown as previously described [30] in 24-well, 35-mm, and 100-mm plates in growth medium (GM) consisting of Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% (v/v) heat-inactivated fetal bovine serum, 1% (v/v) penicillin/streptomycin, and 1% (v/v) L-glutamine at 37 1C in humidified 95% air:5% CO2 atmosphere. For differentiation of myotubes, myoblasts were plated in 0.1% gelatin-coated plates and were grown to 90% confluence. At this point GM was replaced by differentiation medium (DM) consisting of DMEM supplemented with 2% (v/v) heat-inactivated horse serum, 1% (v/v) penicillin/ streptomycin, and 1% (v/v) L-glutamine. During differentiation, DM
191
was replaced every 48 h for 6 days until cell fusion and formation of multinucleated myotubes were achieved. Successful cell differentiation was determined by expression of the main contractile protein MyHC as measured by immunoblotting. Cell media and chemicals were purchased from Biological Industries (Bet HaEmek, Israel). Cell treatments Experiments were conducted on day 7 of differentiation when the cells have completed their differentiation into elongated multinucleated myotubes. Myotubes were exposed to various levels of acetaldehyde or acrolein (Sigma–Aldrich, St. Louis, MO, USA) freshly prepared in DM to final concentrations of 10–500 mM. In each experiment, medium was replaced by fresh DM containing acetaldehyde or acrolein. Control myotubes were treated with fresh DM without acetaldehyde or acrolein. Myotubes were then incubated at 37 1C for various time intervals. In some experiments, myotubes were pretreated with the antioxidant N-acetylcysteine (NAC) or with SB203580, a specific inhibitor of p38 MAPK (Sigma– Aldrich). Myotubes were treated with NAC (2 mM) or SB203580 (5 mM) prepared in fresh DM and incubated for 1 h or 15 min, respectively. Then, medium was replaced by DM containing acetaldehyde or acrolein and myotubes were transferred for incubation at 37 1C. NAC at concentrations of 0.1–30 mM was previously demonstrated to have antioxidant properties in various in vitro systems [32]. Clinically, peak plasma concentration of NAC reached 3.4 mM after intravenous administration of NAC at 150 mg/kg [32]. Estimation of cell viability To assess the effects of acetaldehyde and acrolein on viability of myotubes, 2 105 cells were seeded in 24-well plates and grown to 90% confluence for differentiation to myotubes. On day 7 of differentiation, myotubes were treated with DM containing acetaldehyde or acrolein and incubated for increasing time points (1, 2, 6, 24 h). After treatment, the viability of myotubes was determined by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay (5 mg/ml; Sigma–Aldrich) as previously described [30]. The OD was spectrophotometrically measured in triplicate for each experiment at 570 nm using an ELISA reader (Biochrom Anthos Zenyth, Cambridge, UK). Viability was expressed as a percentage of the values of control myotubes (corresponding to 100%). Estimation of ROS generation To assess the effects of acetaldehyde and acrolein on generation of ROS, 5 105 cells were seeded in 35-mm plates and grown to 90% confluence for differentiation to myotubes. On day 7 of differentiation, myotubes were treated with acetaldehyde or acrolein. After treatment, ROS generation was measured by 2′,7′dichlorofluorescein (DCF) assay as previously described [30]. Briefly, DNA of nuclei was stained with 1.5 mg/ml Hoechst (Invitrogen, Carlsbad, CA, USA) 90 min before treatment. After 30 min, myotubes were washed with phosphate-buffered saline (PBS), loaded with 10 mM CM-H2DCFDA (Invitrogen), and incubated for 1 h. Subsequently, myotubes were washed with PBS and exposed to acetaldehyde or acrolein followed by 20 min of incubation at 37 1C. Then, the myotubes were examined with an Axiovert 135 fluorescence inverted microscope (Carl Zeiss, Oberkochen, Germany) and the fluorescence intensity of DCF was measured using ImageJ software (National Institutes of Health (NIH), Bethesda, MD, USA). Fluorescence intensity was expressed as a percentage of the values of control myotubes (corresponding to 100%).
192
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
Cell lysates and Western blot analysis
Measurement of myotube diameter
After treatments with acetaldehyde or acrolein, myotubes were lysed for cytosolic proteins as previously described [30]. Myotubes were washed twice with PBS and lysed using lysis buffer (50 mM Tris– HCl, pH 7.4, 300 mM NaCl, 1.5 mM MgCl2, 200 mM EDTA, 0.1% Triton X-100) and 40 diluted protease and phosphatase inhibitors (Sigma– Aldrich) and centrifuged at 14,000 rpm for 10 min. Supernatants containing cytosolic proteins were collected and kept at −80 1C. Cytosolic proteins (20 μg) were loaded in each lane and separated by SDS–PAGE. Next, the proteins were transferred to nitrocellulose membranes. Membranes were blocked with 5% nonfat milk powder in Tris-buffered saline–0.125% Tween (TBS-T; Sigma–Aldrich) for 1 h and exposed overnight at 4 1C to primary antibodies. Primary antibodies against the following proteins were used: MyHC (1:1000), MAFbx/atrogin-1 (1:1000), MuRF1 (1:1000) (Santa Cruz Biotechnology, Santa Cruz, CA, USA); actin (1:4000) (Millipore, Billerica, MA, USA); p38 MAPK (1:1000), phospho-p38 MAPK (1:1000) (R&D Systems, Minneapolis, MN, USA). The next day, membranes were washed with TBS-T followed by 1 h incubation at ambient temperature with appropriate secondary antibodies (Jackson ImmunoResearch, West Grove, PA, USA). Detection was performed by enzyme-linked chemiluminescence (ECL; Biological Industries) using an ImageQuant LAS 4000 digital imager system (GE Healthcare, Chalfont St. Giles, Buckinghamshire, UK). Protein quantities were determined by densitometry and analyzed using Total Lab Software V2006C (Nonlinear Dynamics, Newcastle-on-Tyne, UK).
To assess the effects of acetaldehyde and acrolein on cell atrophy, diameter of myotubes was measured after treatments. Myotubes were examined using a phase-contrast microscope (CK40-SLP; Olympus, Tokyo, Japan; objective 20) and digital camera (UC30 Olympus, Japan). Three fields of view from three independent experiments were chosen randomly and the 10 largest myotubes in each field were measured in a blinded fashion using ImageJ software (NIH). Results were expressed as a percentage of the values of control myotubes (corresponding to 100%).
Protein loading control by Ponceau S staining In this study, the effects of acrolein and acetaldehyde on the main contractile muscle proteins MyHC and actin were studied. Therefore, actin could not be used as a housekeeping protein for loading control. As an alternative, quantification of total proteins by reversible Ponceau staining was used for protein loading control as previously described [30]. Briefly, before antibody probing, membranes were rinsed in Ponceau S solution (Bio-Rad, Hercules, CA, USA) for 10 min, followed by a brief rinse in double-distilled water until bands were clearly visible. In each lane, total protein quantities were determined by densitometry and used for normalization of ECL-detected proteins. Quantification of Ponceau staining before antibody probing has been previously validated as an alternative to actin blotting [33]. Real-time PCR RNA purification from myotubes was performed using a High Pure RNA isolation kit (Roche, Mannheim, Germany) according to the manufacturer's instructions. RNA concentrations were quantified at 260 nm by a Nanodrop spectrophotometer (Nanodrop Technologies, Wilmington, DE, USA). Samples of 1 μg RNA were used to synthesize cDNA using a High Capacity cDNA reverse transcription kit (Applied Biosystems, Foster City, CA, USA). QPCR was performed as previously described [30] using a qPCR SYBR Green ROX mix (Thermo Scientific, Basingstoke, UK) and Corbett Rotor-Gene 6000 (Qiagen, Hilden, Germany). For each sample, a value of the threshold cycle (Ct) was calculated using Rotor Gene 6000 series software (Qiagen) based on the time changes in mRNA expression level calculated subsequent to normalization with glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The abundance of target mRNA relative to GAPDH was determined by the ΔΔCt relative quantification method. The following primers were used (Sigma–Aldrich): GAPDH forward, 5′-AGGTCGGTGTGAACGGATTTG-3′, reverse, 5′-TGTAGACCATGTAGTTGAGGTCA-3′; MAFbx/atrogin-1 forward, 5′-CAGCTTCGTGAGCGACCTC-3′, reverse, 5′-GCAGTCGAGAAGTCCAGTC-3′; and MuRF1 forward, 5′-GTGTGAGGTGCCTACTTGCTC-3′, reverse, 5′-GCTCAGTCTTCTGTCCTTGGA-3′.
Statistical analysis Results were expressed as means + standard error (SE) of three independent experiments. Student's t test and one-way ANOVA followed by Tukey's or Dunnett's tests were used for statistical analysis using SPSS 16 software (IBM, USA). A p o 0.05 was considered statistically significant.
Results The effects of acetaldehyde on viability of myotubes To examine the effects of acetaldehyde on catabolism of C2 myotubes, it was essential to assess its effects on viability of myotubes and to find working concentrations at which acetaldehyde would not be cytotoxic. Therefore, differentiated myotubes were exposed to increasing levels of acetaldehyde (up to 500 mM) and incubated at 37 1C. After 24 h, viability of the myotubes was assessed by MTT assay as described under Materials and methods. Exposure of C2 myotubes to acetaldehyde at all concentrations examined did not cause a significant reduction in viability of the myotubes. The effects of acetaldehyde on the viability of myotubes at concentrations of 100, 300, and 500 mM are presented in Fig. 1. The effects of acetaldehyde on ROS generation Recently we have shown that shortly after exposure to CS, generation of ROS in C2 myotubes was increased and later led to muscle catabolism [30]. To assess the role of acetaldehyde in muscle catabolism it was of interest to explore its effects on ROS generation in C2 myotubes. Therefore, differentiated myotubes were exposed to acetaldehyde at increasing levels (up to 500 mM) in the presence or absence of 2 mM NAC and incubated at 37 1C. Twenty minutes after incubation, ROS generation was assessed by
Fig. 1. The effects of acetaldehyde on viability of myotubes. Myotubes were exposed to increasing levels of acetaldehyde and incubated for 24 h. After incubation, viability of the myotubes was measured by MTT assay and compared with untreated myotubes. Results are expressed as means + SE of three different experiments.
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
193
Fig. 2. The effects of acetaldehyde on ROS generation. (A) Untreated myotubes. (B) Myotubes pretreated with vehicle and exposed to 500 mM acetaldehyde. (C) Myotubes pretreated with 2 mM NAC 1 h before exposure to 500 mM acetaldehyde. (D) Changes in fluorescence intensity representing ROS production are expressed as a percentage of the fluorescence intensity found in untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments.
DCF assay as described under Materials and methods. At all concentrations examined, exposure of C2 myotubes to acetaldehyde did not cause a significant increase in ROS production compared with untreated myotubes. Pretreatment with 2 mM NAC before acetaldehyde exposure did not affect ROS generation significantly. The effects of acetaldehyde on ROS generation at the maximal concentration of 500 mM in the presence or absence of NAC are presented in Fig. 2.
Atrogin-1 MuRF1 Acetaldehyde concentration (µM)
CTL
100
300
500
The effects of acetaldehyde on muscle-specific E3 ligases Muscle catabolism is known to be mediated by upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1. To investigate the effects of acetaldehyde on atrogin-1 and MuRF1, C2 myotubes were exposed to acetaldehyde at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After incubation with acetaldehyde, protein levels of atrogin-1 and MuRF1 were examined by Western blot as described under Materials and methods. At all concentrations and time points examined, no significant change in protein levels of atrogin-1 or MuRF1 was found after exposure to acetaldehyde. The effects of 24 h exposure to acetaldehyde on protein levels of atrogin-1 and MuRF1 at concentrations of 100, 300, and 500 mM are presented in Fig. 3. The effects of acetaldehyde on muscle contractile proteins and diameter of myotubes To examine the effects of acetaldehyde on the main contractile muscle proteins MyHC and actin, C2 myotubes were exposed to acetaldehyde at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After incubation with acetaldehyde, protein levels of MyHC and actin were examined by Western blot. At all concentrations and time points examined, no significant change in protein levels of MyHC and actin was found after exposure to acetaldehyde. The effects of 24 h exposure
Fig. 3. The effects of acetaldehyde on muscle-specific E3 ligases. Myotubes were incubated with increasing concentrations of acetaldehyde (up to 500 mM) for 24 h. Untreated myotubes served as control. After incubation, cell lysates were prepared and subjected to Western blot analysis with antibodies against atrogin-1 and MuRF1. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are expressed as means + SE of three independent experiments.
to acetaldehyde on protein levels of MyHC and actin at concentrations of 100, 300, and 500 mM are presented in Fig. 4A. In addition, after incubation with acetaldehyde, myotubes were photographed and diameters were measured and compared with untreated myotubes as described under Materials and methods. Similarly, at all concentrations and time points examined, no significant change in the diameter of myotubes was found after exposure to acetaldehyde. Representative images of myotubes exposed to the
194
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
MyHC Actin Acetaldehyde concentration (µM)
CTL
100
300
500
Fig. 4. The effects of acetaldehyde on muscle contractile proteins and diameter of myotubes. (A) Myotubes were incubated with increasing concentrations of acetaldehyde (up to 500 mM) for 24 h. Untreated myotubes served as control. After incubation, cell lysates were prepared and subjected to Western blot analysis with antibodies against MyHC and actin. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are expressed as means + SE of three independent experiments. (B) Untreated myotubes. (C) Myotubes exposed to 500 mM acetaldehyde for 24 h. (D) Changes in diameter of myotubes are expressed as percentage of the diameter of untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments.
maximal concentration of 500 mM for 24 h in comparison with untreated myotubes are presented in Fig. 4B–D.
The effects of acrolein on viability of myotubes Acrolein is known as an extremely toxic unsaturated aldehyde [7]. Therefore, examining the effects of acrolein on the viability of C2 myotubes was essential to find a suitable concentration for experiments of acrolein exposure. Thus, myotubes were exposed to acrolein at increasing concentrations (up to 500 mM) and for increasing time periods (up to 24 h). After exposure, the viability
of myotubes was assessed by MTT assay as described under Materials and methods. After 1 h incubation, exposure to acrolein at concentrations up 100 mM did not cause a significant reduction in cell viability. However, exposure to acrolein at 250 and 500 mM caused a significant reduction in the viability of myotubes (Fig. 5A). Subsequently, the effects of 25 mM acrolein on the viability of myotubes at increasing incubation periods were examined. After 6 h of incubation with 25 mM acrolein, the viability of myotubes decreased significantly. However, at 6 h, the viability of myotubes remained higher than 80% (Fig. 5B). Therefore, experiments were carried out with acrolein at a concentration of 25 mM for up to 6 h.
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
195
Acrolein increases ROS generation in myotubes Myotubes were exposed to 25 mM acrolein in the presence or absence of 2 mM NAC and incubated at 37 1C. After 20 min, ROS generation was assessed by the DCF assay. Exposure of C2 myotubes to acrolein caused a significant increase in ROS generation in comparison with untreated myotubes. Pretreatment with 2 mM NAC 1 h before acrolein exposure prevented acrolein-induced increase in ROS generation. Representative images of myotubes exposed to 25 mM acrolein in the presence or absence of NAC and compared with untreated myotubes are presented in Fig. 6. Acrolein upregulates muscle-specific E3 ligases To investigate the effects of acrolein on the expression of atrogin-1 and MuRF1, C2 myotubes were exposed to 25 mM acrolein and incubated for increasing time periods (up to 6 h). After incubation, protein levels of atrogin-1 and MuRF1 were examined by Western blot and mRNA levels were determined by qPCR as described under Materials and methods. A significant increase in mRNA levels of atrogin-1 and MuRF1 compared with untreated myotubes was detected at 1 and 2 h of incubation with acrolein (Fig. 7A). In addition, a significant increase in protein levels of atrogin-1 and MuRF1 was detected from 1 h until 6 h of incubation with acrolein (Fig. 7B). Acrolein induces MyHC degradation and reduction in diameter of myotubes
Fig. 5. The effects of acrolein on viability of myotubes. (A) Myotubes were exposed to increasing levels of acrolein and incubated for 1 h. (B) Myotubes were exposed to acrolein at 25 mM for increasing incubation periods. After incubation, the viability of myotubes was measured by MTT assay and compared with that of untreated myotubes. Results are expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
To examine the effects of acrolein on the atrophy of myotubes and the main contractile muscle proteins MyHC and actin, myotubes were exposed to 25 mM acrolein and incubated for increasing time intervals (up to 6 h). After incubation, myotubes were photographed and diameters were measured and compared with those of untreated myotubes. Subsequently, myotubes were lysed
Fig. 6. Acrolein increases ROS generation in myotubes. (A) Untreated myotubes. (B) Myotubes pretreated with vehicle and exposed to 25 mM acrolein. (C) Myotubes pretreated with 2 mM NAC 1 h before exposure to 25 mM acrolein. (D) Changes in fluorescence intensity representing ROS production are expressed as percentage of the fluorescence intensity found in untreated myotubes. Results are relative to control and expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
196
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
myotubes were pretreated with 5 μM p38 MAPK inhibitor SB203580 or vehicle 15 min before exposure to 25 mM acrolein and incubation (up to 6 h). After incubation, phosphorylation of p38 MAPK and protein levels of atrogin-1, MuRF1, MyHC, and actin were examined by Western blot. Significant phosphorylation of p38 MAPK was observed at 1 and 2 h of exposure to acrolein. Pretreatment with SB203580 blocked acrolein-induced activation of p38 MAPK (Fig. 9A). In addition, inhibition of p38 MAPK abolished acrolein-induced increase in protein levels of atrogin-1 and MuRF1 and degradation of MyHC at 2 h of exposure to acrolein. Representative blots are presented in Fig. 9B and C.
Discussion
Atrogin -1 MuRF1 Acrolein (25 µM) incubation time
C T L 1 hr
2 hr
6 hr
Fig. 7. Acrolein upregulates muscle-specific E3 ligases. Myotubes were exposed to 25 mM acrolein and incubated for increasing time periods. Untreated myotubes served as control. (A) After incubation, RNA was isolated and subjected to reverse transcription and qPCR analysis to determine the expression of atrogin-1 and MuRF1. Data were normalized to GAPDH expression and are relative to the corresponding value of control myotubes. (B) After incubation with acrolein, cell lysates were prepared and subjected to Western blot analysis with antibodies against atrogin-1 and MuRF1. Protein levels were normalized to total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control myotubes. Results are expressed as means + SE of three independent experiments. *p o 0.05 versus control myotubes.
and subjected to Western blot analysis. Diameter of myotubes decreased depending on acrolein incubation time. A significant decrease in diameter was found at 2 and 6 h of incubation with acrolein compared with untreated myotubes. Representative images are presented in Fig. 8A–E. Also, protein levels of MyHC decreased depending on acrolein incubation times. A significant decrease in MyHC was found at 2 and 6 h of incubation with acrolein compared with untreated myotubes. The protein level of actin did not change significantly. Representative blots are presented in Fig. 8F. The involvement of p38 MAPK in acrolein-induced muscle catabolism In previous studies, activation of p38 MAPK was shown to be one of the initial steps for CS-induced catabolism of skeletal muscle [14,30,34]. Moreover, inhibition of p38 MAPK abolished the catabolic effects caused by exposure to CS [30] or extract of CS [34]. Therefore, it was of interest to examine the role of p38 MAPK in acrolein-induced catabolism of myotubes. Thus,
In this study, skeletal myotubes were exposed to various levels of acetaldehyde and acrolein to assess their role in catabolism of skeletal muscle. For the first time, it was shown that exposure of skeletal myotubes to acrolein and not acetaldehyde resulted in cell atrophy and breakdown of MyHC through increased oxidative stress, activation of p38 MAPK, and upregulation of the musclespecific E3 ligases atrogin-1 and MuRF1. Several conditions in which levels of acetaldehyde and acrolein may be increased were associated with muscle catabolism. Acetaldehyde was previously suggested to damage skeletal muscle in alcoholic myopathy [3,4]. In addition, acetaldehyde is one of the major constituents of CS [19]. Epidemiological, clinical, in vivo, and in vitro studies reported that smoking or exposure to CS can damage skeletal muscle and promote its catabolism [14,25–31,34]. Therefore, it was of great interest to study the effects of acetaldehyde exposure on muscle catabolism in skeletal myotubes. Blood levels of acetaldehyde in healthy subjects were reported at 12.2 mM [35]. After administration of alcohol, mean concentrations of acetaldehyde in alcoholics were 42.7 mM in comparison with 26.5 mM in nonalcoholics [36]. Smoking a single cigarette can elevate blood levels of acetaldehyde by up to 9.5 mM [19]. In this study, the catabolic effects of acetaldehyde were investigated in physiological and supraphysiological levels of up to 500 mM. At all concentrations and time points examined, acetaldehyde was not cytotoxic and did not promote catabolism of skeletal myotubes. C2 myotubes exposed to acetaldehyde in concentrations of up to 500 mM did not demonstrate increased ROS generation or upregulation of atrogin-1 and MuRF1. Also, no significant reductions in the diameter of myotubes or in the levels of the main contractile muscle proteins MyHC and actin were found. It was previously shown that exposure of human myocytes to 200 mM acetaldehyde decreased basal protein synthesis without altering the rate of protein degradation [5]. Also, injection of acetaldehyde into rats resulted in reduced fractional rates of protein synthesis in skeletal muscles [6]. Therefore, it is possible that acetaldehyde may promote catabolism of skeletal muscle, not through the activation of catabolic pathways but through inhibition of anabolic pathways. To prove this hypothesis, future studies should focus on the effects of acetaldehyde on inhibition of anabolic pathways in skeletal muscle. These pathways include the insulin-like growth factor 1/phosphatidylinositol 3-kinase/protein kinase B and the mammalian target of rapamycin signaling pathways, which regulate protein synthesis in skeletal muscle [37]. Acrolein has numerous biological effects including regulation of signaling molecules and activation of transcription factors [7]. Conditions in which blood or tissue levels of acrolein may be increased include renal failure, oxidative stress, and heavy smoking [10,11]. During inflammation and oxidative stress acrolein can be produced endogenously via myeloperoxidase-mediated degradation of threonine and amine oxidase-mediated degradation of spermine and spermidine [9]. Wasting of skeletal muscle is
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
197
MyHC Actin Acrolein (25 µM) incubation time
CTL
1 hr
2 hr
6 hr
Fig. 8. Acrolein induces MyHC degradation and reduction in diameter of myotubes. (A) Untreated myotubes. (B) Myotubes exposed to 25 mM acrolein for 1 h. (C) Myotubes exposed to 25 mM acrolein for 2 h. (D) Myotubes exposed to 25 mM acrolein for 6 h. (E) Changes in diameter of myotubes are expressed as a percentage of the diameter of untreated myotubes. (F) Cell lysates were prepared and subjected to Western blot analysis with antibodies against MyHC and actin. Protein levels were normalized by total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control. Results are relative to control and expressed as means + SE of three different experiments. *p o 0.05 versus control myotubes.
common in renal failure patients [13] and may be increased in situations of oxidative stress and chronic inflammation [14]. The main source of exogenous acrolein in humans is exposure to CS [2,9]. CS is considered to make up a large proportion of total human exposure to acrolein [9]. As mentioned earlier, the catabolic effects of CS were demonstrated previously [14,25–31,34]. Therefore, it was of great interest to assess the effects of acrolein exposure on catabolism of skeletal myotubes. To the best of our knowledge, the effects of acrolein exposure on skeletal muscle have not been studied previously. However, the effects of acrolein on cardiac muscle were investigated earlier. Ismahil et al. [15] found that chronic oral exposure of mice to acrolein (1 mg/kg) resulted in cardiomyopathy accompanied by contractile dysfunction and myocardial oxidative stress. Also, Luo et al. [1] showed that intravenous injection of acrolein (0.5 mg/kg) into mice resulted in myocardial dysfunction and increased acrolein–protein adducts, including adducts to sarcomeric proteins such as myosin light polypeptide. The above studies demonstrate that acrolein administered both orally and intravenously can reach cardiac muscle tissue and promote cardiomyopathy. Previous studies investigated the effects of acrolein on smooth muscle cultures and demonstrated its role in activation of signaling pathways and gene regulation. Ranganna et al. [16] found that exposure of rat vascular smooth muscle cells to acrolein resulted in activation of several members of the MAPK family, including p38 MAPK. A concentration- and time-dependent activation of p38 MAPK was induced by acrolein and was inhibited by SB203580 pretreatment. Volpi et al. [17] exposed human airway smooth muscle cells to CS extract or acrolein at concentrations of 10–100 mM. They found that acrolein mimicked the effects of CS
extract and caused activation of p38 MAPK, which led to upregulation of vascular endothelial growth factor. Acrolein at 60 mM elicited a rapid phosphorylation of p38 MAPK that peaked at 1 h and was blocked by SB203580. Similarly, Moretto et al. [18] exposed human bronchial smooth muscle cells to CS extract or acrolein at concentrations of 10–60 mM. Both CS extract and acrolein induced p38 MAPK phosphorylation, which resulted in upregulation of interleukin-8. Peak phosphorylation was found at 30 min of exposure to 30 mM acrolein and was abolished by SB203580. In this study, it is shown that exposure of C2 myotubes to acrolein at the level of 25 mM caused atrophy of myotubes and breakdown of MyHC via increased ROS generation, activation of p38 MAPK, and upregulation of atrogin-1 and MuRF1. Similar to the above studies, peak phosphorylation of p38 MAPK was observed at 1 h of exposure to acrolein and was prevented by SB203580 pretreatment. Recently, we have shown that CS exposure to skeletal myotubes also caused cell atrophy and degradation of MyHC through increased oxidative stress, activation of p38 MAPK, and upregulation of atrogin-1 and MuRF1. Inhibition of p38 MAPK prevented CS-induced muscle catabolism in skeletal myotubes [30]. In this study, acrolein was shown to mimic the effects of CS by activating a similar signaling cascade that resulted in muscle catabolism. Because acrolein mimicked the effects of CS on skeletal muscle, it may be suggested that acrolein is one of the main components of CS responsible for its catabolic effects. However, CS is a complex aerosol consisting of thousands of constituents [38]. Various constituents of CS have the potential to induce catabolism of skeletal muscle [31]. For instance, CS is a major source of both ROS
198
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
P-p38 p38 Acrolein (25 µM) CTL incubation time
1 hr
1 hr
2 hr
2 hr
6 hr
6 hr
SB203580
-
+
-
+
-
+
-
Atrogin-1
MyHC
MuRF1
Actin
Acrolein (25 µM) incubation time SB203580
CTL
2 hr
2 hr
-
-
+
Acrolein (25 µM) incubation time SB203580
CTL
2 hr
2 hr
-
-
+
Fig. 9. The involvement of p38 MAPK in acrolein-induced muscle catabolism. Myotubes were exposed to 25 mM acrolein in the presence of 5 μM SB203580 or vehicle. Untreated myotubes served as control. After incubation with acrolein, cell lysates were prepared and subjected to Western blot analysis with antibodies against p38 MAPK (p38), phosphorylated p38 MAPK (P-p38), atrogin-1, MuRF1, MyHC, and actin. (A) Protein levels of P-p38 and p38 were quantified by densitometry and values of P-p38 were normalized to p38 and compared with control. (B and C) Protein levels were normalized by total protein densitometry detected by Ponceau S staining and expressed relative to the corresponding value of control myotubes. Results are expressed as means + SE of three independent experiments. *p o 0.05 versus control myotubes.
and reactive nitrogen species (RNS) [31]. ROS present in CS, such as hydrogen peroxide (H2O2), were shown to upregulate atrogin-1 and MuRF1 and stimulate ubiquitination of muscle proteins in skeletal myotubes [39]. Also, RNS present in CS, such as peroxynitrite (ONOO−), were demonstrated to cause degradation of muscle proteins in cultured myotubes mediated by activation of nuclear factor-κB [40]. Therefore, it seems that in addition to acrolein, ROS and RNS from CS may also play an important role in the catabolism of skeletal muscle in smokers. Our findings demonstrate that acrolein-induced catabolism of skeletal muscle is related to increased oxidative stress as found by the DCF assay (Fig. 6). Acrolein may increase ROS production and oxidative stress in several mechanisms. First, acrolein can activate the nicotinamide adenine dinucleotide phosphate oxidase complex, leading to superoxide production. Superoxide dismutase converts superoxide into H2O2 and activates inducible nitric oxide synthase, leading to production of nitric oxide and formation of ONOO−, which ultimately causes oxidative and nitrosative stress [41]. In addition, acrolein can react with cellular nucleophiles, such as reduced glutathione (GSH) [41]. The conjugation of acrolein with GSH depletes its cellular levels, impairing the clearance of free radicals that results in increased oxidative stress [16,41].
The mechanisms by which acrolein increases oxidative stress in skeletal muscle should be investigated in future studies. This study has certain limitations that need to be addressed when comparing in vitro exposure of acrolein with clinical conditions of increased exposure to acrolein. First, myotubes were treated with acrolein prepared in low-protein medium consisting of 2% horse serum as required for their successful differentiation from myoblasts into myotubes. However, human plasma contains a higher percentage of protein (approximately 6–8%). Thus, a higher percentage of protein in the medium as in plasma might have a different impact on the cytotoxicity and biological effects of acrolein that were found in this study. In addition, the working concentration of acrolein (25 mM) was determined from viability assays that examined the effects of acrolein on the viability of myotubes at increasing concentrations of up to 500 mM. Another limitation is the difficulty in comparing the concentrations of acrolein used in our in vitro study with its physiological levels in various conditions. For instance, during inflammation, phagocytes can generate acrolein by the action of myeloperoxidase on threonine that can circulate in high concentrations of about 2 mM [10]. In renal failure patients, plasma levels of acrolein were studied previously. Using high-performance
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
liquid chromatography (HPLC), Igarashi et al. [12] found that free acrolein in plasma of uremic patients was 1.4 mM, in comparison with 0.5 mM in control subjects. Acrolein found as protein conjugates in uremic patients was equivalent to 170 mM, about fivefold higher than its concentration in the plasma of control subjects. Thus, in comparison with free acrolein levels found in plasma of renal failure patients, the acrolein concentration of 25 mM used in this study appears to be supraphysiological. In heavy smokers, circulating or tissue concentrations of acrolein are increased [10,11]. However, as far as we are aware, the exact concentrations of acrolein in the plasma of smokers and the levels to which skeletal muscles of smokers are exposed to acrolein have not been studied yet. Annovazzi et al. [23] performed HPLC analysis on the saliva of smokers versus nonsmokers and showed that average acrolein concentrations in heavy smokers were 21 mM compared with 2.3 mM in nonsmokers. Eiserich et al. [21] calculated that the concentration of acrolein in respiratory tract lining fluids (RTLF) after exposure to one cigarette is 80 mM. Carmella et al. [24] found that the levels of 3,3-hydroxypropylmercapturic acid, a major acrolein metabolite formed by GSH conjugation, in urine are related to cigarette smoking, suggesting that acrolein is transported from the lung into the systemic circulation. The calculation by Eiserich et al. [21] suggests that the acrolein concentration of 25 mM used in this study may be within the physiological range. However, this calculation was based on the assumption that all of the components of CS are completely deposited within the RTLF with no further metabolism. Thus, RTLF and plasma concentrations of acrolein after exposure to one cigarette may be lower than calculated. On the other hand, the above calculation was based on experiments that examined exposure to a single cigarette without considering the effects of prolonged exposure to acrolein as in heavy smokers. Therefore, until the exact level of acrolein to which skeletal muscles of smokers are exposed is investigated, it will be difficult to compare the concentrations of acrolein used in our in vitro study with its physiological levels in smokers.
Conclusions Our findings demonstrate that exposure of skeletal myotubes to acrolein and not acetaldehyde resulted in atrophy of myotubes and breakdown of MyHC via increased oxidative stress, activation of p38 MAPK, and upregulation of the muscle-specific E3 ligases atrogin-1 and MuRF1. Although within the limitations of an in vitro study, these findings may provide a molecular explanation for muscle wasting in conditions of increased exposure to acrolein, including cigarette smoking, renal failure, oxidative stress, and inflammation. Future clinical studies should examine the role of acrolein in wasting and dysfunction of skeletal muscle in these pathologies and in heavy smokers.
Acknowledgments This study was supported by grants from the Rappaport Institute, the Krol Foundation of Barnegat, New Jersey, the Myers–JDC Brookdale Institute of Gerontology and Human Development, and ESHEL—the association for planning and development of services for the aged in Israel. Special thanks to Dr. Alvira Bromosov for her help in the DCF assay.
199
References [1] Luo, J.; Hill, B. G.; Gu, Y.; Cai, J.; Srivastava, S.; Bhatnagar, A.; Prabhu, S. D. Mechanisms of acrolein-induced myocardial dysfunction: implications for environmental and endogenous aldehyde exposure. Am. J. Physiol. Heart Circ. Physiol 293:H3673–H3684; 2007. [2] Voulgaridou, G. P.; Anestopoulos, I.; Franco, R.; Panayiotidis, M. I.; Pappa, A. DNA damage induced by endogenous aldehydes: current state of knowledge. Mutat. Res. 711:13–27; 2011. [3] Preedy, V. R.; Keating, J. W.; Peters, T. J. The acute effects of ethanol and acetaldehyde on rates of protein synthesis in type I and type II fiber-rich skeletal muscles of the rat. Alcohol Alcohol 27:241–251; 1992. [4] Preedy, V. R.; Adachi, J.; Ueno, Y.; Ahmed, S.; Mantle, D.; Mullatti, N.; Rajendram, R.; Peters, T. J. Alcoholic skeletal muscle myopathy: definitions, features, contribution of neuropathy, impact and diagnosis. Eur. J. Neurol. 8:677–687; 2001. [5] Hong-Brown, L. Q.; Frost, R. A.; Lang, C. H. Alcohol impairs protein synthesis and degradation in cultured skeletal muscle cells. Alcohol Clin. Exp. Res. 25:1373–1382; 2001. [6] Oba, T.; Maeno, Y.; Ishida, K. Differential contribution of clinical amounts of acetaldehyde to skeletal and cardiac muscle dysfunction in alcoholic myopathy. Curr. Pharm. Des 11; 2005. 791-780. [7] Berry, K. A.; Henson, P. M.; Murphy, R. C. Effects of acrolein on leukotriene biosynthesis in human neutrophils. Chem. Res. Toxicol. 21:2424–2432; 2008. [8] Anderson, E. J.; Katunga, L. A.; Willis, M. S. Mitochondria as a source and target of lipid peroxidation products in healthy and diseased heart. Clin. Exp. Pharmacol. Physiol. 39:179–193; 2012. [9] Stevens, J. F.; Maier, C. S. Acrolein: sources, metabolism, and biomolecular interactions relevant to human health and disease. Mol. Nutr. Food Res. 52:7–25; 2008. [10] Gugliucci, A. Antithrombin activity is inhibited by acrolein and homocysteine thiolactone: protection by cysteine. Life Sci 82:413–418; 2008. [11] Misonou, Y.; Asahi, M.; Yokoe, S.; Miyoshi, E.; Taniguchi, N. Acrolein produces nitric oxide through the elevation of intracellular calcium levels to induce apoptosis in human umbilical vein endothelial cells: implications for smoke angiopathy. Nitric Oxide 14:180–187; 2006. [12] Igarashi, K.; Ueda, S.; Yoshida, K.; Kashiwagi, K. Polyamines in renal failure. Amino Acids 31:477–483; 2006. [13] Sukhanov, S.; Semprun-Prieto, L.; Yoshida, T.; Tabony, A. M.; Higashi, Y.; Galvez, S.; Delafontaine, P. Angiotensin II, oxidative stress and skeletal muscle wasting. Am. J. Med. Sci. 342:143–147; 2011. [14] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Sarcopenia and smoking: a possible cellular model of cigarette smoke effects on muscle protein breakdown. Ann. N. Y. Acad. Sci 1259:47–53; 2011. [15] Ismahil, M. A.; Hamid, T.; Haberzettl, P.; Gu, Y.; Chandrasekar, B.; Srivastava, S.; Bhatnagar, A.; Prabhu, S. D. Chronic oral exposure to the aldehyde pollutant acrolein induces dilated cardiomyopathy. Am. J. Physiol. Heart Circ. Physiol 301:2050–2060; 2011. [16] Ranganna, K.; Yousefipour, Z.; Nasif, R.; Yatsu, F. M.; Milton, S. G.; Hayes, B. E. Acrolein activates mitogen-activated protein kinase signal transduction pathways in rat vascular smooth muscle cells. Mol. Cell. Biochem. 240:83–98; 2002. [17] Volpi, G.; Facchinetti, F.; Moretto, N.; Civelli, M.; Patacchini, R. Cigarette smoke and α,β-unsaturated aldehydes elicit VEGF release through the p38 MAPK pathway in human airway smooth muscle cells and lung fibroblasts. Br. J. Pharmacol 163:649–661; 2011. [18] Moretto, N.; Facchinetti, F.; Southworth, T.; Civelli, M.; Singh, D.; Patacchini, R. α,β-Unsaturated aldehydes contained in cigarette smoke elicit IL-8 release in pulmonary cells through mitogen-activated protein kinases. Am. J. Physiol. Lung Cell. Mol. Physiol 296:839–848; 2009. [19] Talhout, R.; Opperhuizen, A.; Van Amsterdam, J. G. Role of acetaldehyde in tobacco smoke addiction. Eur. Neuropsychopharmacol. 17:627–636; 2007. [20] Smith, C. J.; Fischer, T. H. Particulate and vapor phase constituents of cigarette mainstream smoke and risk of myocardial infarction. Atherosclerosis 158: 257–267; 2011. [21] Eiserich, J. P.; Van der Vliet, A.; Handelman, G. J.; Halliwell, B.; Cross, C. E. Dietary antioxidants and cigarette smoke-induced biomolecular damage: a complex interaction. Am. J. Clin. Nutr. 62:1490S–1500S; 1995. [22] Colombo, G.; Aldini, G.; Orioli, M.; Giustarini, D.; Gornati, R.; Rossi, R.; Colombo, R.; Carini, M.; Milzani, A.; Dalle-Donne, I. Water-soluble alpha, beta-unsaturated aldehydes of cigarette smoke induce carbonylation of human serum albumin. Antioxid. Redox Signaling 12:349–364; 2010. [23] Annovazzi, L.; Cattaneo, V.; Viglio, S.; Perani, E.; Zanone, C.; Rota, C.; Pecora, F.; Cetta, G.; Silvestri, M.; Iadarola, P. High-performance liquid chromatography and capillary electrophoresis: methodological challenges for the determination of biologically relevant low-aliphatic aldehydes in human saliva. Electrophoresis 25:1255–1263; 2004. [24] Carmella, S. G.; Chen, M.; Zhang, Y.; Zhang, S.; Hatsukami, D. K.; Hecht, S. S. Quantitation of acrolein-derived (3-hydroxypropyl)mercapturic acid in human urine by liquid chromatography–atmospheric pressure chemical ionization tandem mass spectrometry: effects of cigarette smoking. Chem. Res. Toxicol. 20:986–990; 2007. [25] Petersen, A. M.; Magkos, F.; Atherton, P.; Selby, A.; Smith, K.; Rennie, M. J.; Pedersen, B. K.; Mittendorfer, B. Smoking impairs muscle protein synthesis and increases the expression of myostatin and MAFbx in muscle. Am. J. Physiol. Endocrinol. Metab. 293:843–848; 2007.
200
O. Rom et al. / Free Radical Biology and Medicine 65 (2013) 190–200
[26] Montes de Oca, M.; Loeb, E.; Torres, S. H.; De Sanctis, J.; Hernández, N.; Tálamo, C. Peripheral muscle alterations in non-COPD smokers. Chest 133:13–18; 2008. [27] Castillo, E. M.; Goodman-Gruen, D.; Kritz-Silverstein, D.; Morton, D. J.; Wingard, D. L.; Barrett-Connor, E. Sarcopenia in elderly men and women: the Rancho Bernardo study. Am. J. Prev. Med. 25:226–231; 2003. [28] Szulc, P.; Duboeuf, F.; Marchand, F.; Delmas, P. D. Hormonal and lifestyle determinants of appendicular skeletal muscle mass in men: the MINOS study. Am. J. Clin. Nutr. 80:496–503; 2004. [29] Lee, J. S.; Auyeung, T. W.; Kwok, T.; Lau, E. M.; Leung, P. C.; Woo, J. Associated factors and health impact of sarcopenia in older Chinese men and women: a cross-sectional study. Gerontology 53:404–410; 2007. [30] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Cigarette smoke and muscle catabolism in C2 myotubes. Mech. Ageing Dev. 134; 2013. 24-23. [31] Rom, O.; Kaisari, S.; Aizenbud, D.; Reznick, A. Z. Identification of possible cigarette smoke constituents responsible for muscle catabolism. J. Muscle Res. Cell Motil. 33:199–208; 2012. [32] Sadowska, A. M.; Manuel-Y-Keenoy, B.; De Backer, W. A. Antioxidant and antiinflammatory efficacy of NAC in the treatment of COPD: discordant in vitro and in vivo dose-effects: a review. Pulm. Pharmacol. Ther 20:9–22; 2007. [33] Romero-Calvo, I.; Ocon, B.; Martinez-Moya, P.; Suarez, M. D.; Zarzuelo, A.; Martinez-Augustin, O.; Medina, F. S. Reversible Ponceau staining as a loading control alternative to actin in Western blots. Anal. Biochem. 401:318–320; 2010.
[34] Liu, Q.; Xu, W. G.; Luo, Y.; Han, F. F.; Yao, X. H.; Yang, T. Y.; Zhang, Y.; Pi, W. F.; Guo, X. J. Cigarette smoke-induced skeletal muscle atrophy is associated with up-regulation of USP-19 via p38 and ERK MAPKs. J. Cell. Biochem. 112:2307–2316; 2011. [35] Pezzoli, C.; Galli-kienle, M.; Padova, C.; Stramentinoli, G. HPLC method for the evaluation of blood acetaldehyde without ethanol interference. J. Liq. Chromatogr 7:765–778; 1984. [36] Korsten, M. A.; Matsuzaki, S.; Feinman, L.; Lieber, C. S. High blood acetaldehyde levels after ethanol administration: difference between alcoholic and nonalcoholic subjects. N. Engl. J. Med. 292:386–389; 1975. [37] Meng, S. J.; Yu, L. J. Oxidative stress, molecular inflammation and sarcopenia. Int. J. Mol. Sci. 11:1509–1526; 2010. [38] Swan, E. G.; Lessov-Schlaggar, C. N. The effects of tobacco smoke and nicotine on cognition and the brain. Neuropsychol. Rev. 17:259–273; 2007. [39] Li, Y.; Chen, Y.; Li, A. S.; Reid, M. B. Hydrogen peroxide stimulates ubiquitinconjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes. Am. J. Physiol. Cell Physiol 285:C806–C812; 2003. [40] Bar-Shai, M.; Reznick, A. Z. Reactive nitrogen species induce nuclear factor-κBmediated protein degradation in skeletal muscle cells. Free Radic. Biol. Med 40:2112–2125; 2006. [41] Ansari, M. A.; Keller, J. N.; Scheff, S. W. Protective effect of Pycnogenol in human neuroblastoma SH-SY5Y cells following acrolein-induced cytotoxicity. Free Radic. Biol. Med. 45:1510–1519; 2008.