Salmeterol is a β2-adrenergic receptor agonist that is commonly used in the treatment of asthma and chronic obstructive pulmonary disease (COPD). β2-adrenergic receptor agonists are divided into two main groups according to their molecular properties. The short-acting β-adrenergic agonists (e.g., salbutamol) are used regularly for asthma flares, acutely for rescue bronchodilation after exposure to an allergen or irritant, or in preparation for exercise in the case of exercise-induced bronchospasm (23,35). The long-acting β-adrenergic agonists (e.g., formeterol and salmeterol) are used for moderate-to-severe persistent asthma, moderate-to-severe COPD, maintenance of bronchodilation, control of bronchospasms, and control of nocturnal symptoms in asthma and other obstructive diseases.
Salmeterol is >10,000 times more lipophilic than salbutamol. The use of low-angle neutron diffraction techniques has shown that salmeterol diffuses rapidly into the cell membrane and then moves laterally to approach the active site of the β2-adrenoceptor through the membrane (21). Molecular biology studies have shown that a discrete portion of the fourth transmembrane domain of the β2-adrenoceptor, believed to be associated with exosite binding, is essential for the long-lasting activity of salmeterol. The mechanism of action of salmeterol therefore involves the interaction of the side chain with an auxiliary binding site (exosite), a motif composed of highly hydrophobic amino acids within the fourth domain of the β2-adrenoceptor. When the side chain is associated with the exosite, the molecule is prevented from dissociating from the β2-adrenoceptor. However, the head can still freely engage and disengage the active site by the Charniére (hinge) principle, with the fulcrum being the oxygen atom in the side chain. The position of this oxygen atom is critical for this long-acting drug (21). The duration of action of β2 agonists in the human bronchus is in the following order: salmeterol >> formoterol ≥ salbutamol ≥ terbutaline > fenoterol (21). Long-acting β2 agonists provide more sustained bronchodilation for 12 h, reduce daytime and nighttime symptoms, and improve the quality of life in adult asthmatic patients better than short-acting β2 agonists.
Different types of skeletal muscle fibers express different levels and types of the β-adrenergic receptor (26). Recent work has focused on characterizing the function of β-adrenergic receptors in skeletal muscle, but their functional roles have not been completely elucidated. These receptors are likely involved in several aspects of muscle function including the stimulation of glycogenolysis, triglyceride lipolysis, oxygen consumption in the muscle, and ion exchange. They can also promote muscle satellite cell activation (34), thereby increasing force generation in the muscle. These effects could differ in various muscles due to the presence of different numbers and/or types of β-adrenergic receptors. Previous studies have demonstrated that chronic administration of β-adrenoceptor (β-AR) agonists (particularly β2-AR agonists) can increase myofibrillar protein content and induce skeletal muscle hypertrophy in mammals (30). This β2-AR–induced hypertrophy is believed to be a result of decreased proteolysis coupled with increased protein synthesis. The ubiquitin–proteasome signaling cascade, Ca2+-dependent proteolysis, and/or calpain-mediated proteolysis have all been proposed to play a role; however, the molecular and cellular pathways altered after β-AR agonist administration remain poorly understood (24). In addition to hypertrophy, the acute exposure of skeletal muscle tissue and cells to β-AR agonists has been found to modulate oxidative metabolism, energy expenditure, lipolysis, glucose transport, and glucose oxidation (19,28).
In sport practice, β2-AR agonists are used by asthmatic individuals to counteract asthmatic episodes or to prevent exercise-induced asthma and pulmonary edema, mainly in winter sports (39). However, because of their ability to ameliorate bronchodilation, promote muscle hypertrophy, and induce protein turnover, competitive and recreational athletes and body builders have often used β2-AR agonists as performance-enhancing agents (8,11). Although muscle hypertrophy is not strictly correlated with increased sport performance, the illicit use of these agents persists (9). The extensive animal-based research makes it clear that the administration of β-agonists (especially in high doses) can produce significant undesirable physiological adverse effects that are likely to deleteriously affect athletic performance in the long term. The most frequently adverse effects associated with the use of salmeterol include nausea, headaches, muscle tremor, palpitation, muscle cramp, peripheral vasodilatation, tachycardia, and myocardial infarction (29). Moreover, it has been demonstrated that the β2-AR agonist clenbuterol is capable of inducing myocyte apoptosis (3). Furthermore, the onset of myocyte death can occur at doses lower than those needed to induce muscle hypertrophy (43). It is therefore important to ascertain whether this class of molecules can induce apoptosis in striated muscles, similar to the well-known toxic effects of catecholamines.
Although the presence of numerous studies on β2-AR agonists, the cellular effects of high concentrations of salmeterol in skeletal muscle have been not investigated yet. To clarify the effects of this molecule in skeletal muscle precursor and muscle postmitotic tissue, C2C12 (mouse) and L6C5 (rat) myogenic cells lines were used in proliferative (myoblasts) or differentiated (myotubes) state to mimic the responses of satellite cells and muscle fibers, respectively.
In this study, we demonstrate that exposure of proliferating and differentiated cells to salmeterol shows a different effect depending on treatment conditions. While short-term treatment with high supratherapeutic doses of the drug increases cell metabolism without affecting cell growth and viability, prolonged exposure leads to cell growth arrest and cell death.
Indeed, our results show for the first time that salmeterol treatment at supratherapeutic doses, miming doping abuse, induces skeletal muscle cell death by activation of the intrinsic apoptotic pathway.
Cell Culture and Materials
Mouse C2C12 (ATCC, Manassas, VA) and rat L6C5 (a clone derived from the L6 muscle cell line; ICLC, Genova, Italy) myoblasts were grown in Dulbecco modified Eagle medium supplemented with Glutamax-I (L-alanyl-L-glutamine), 4500 mg·L−1 glucose (Invitrogen, Carlsbad, CA), and 10% (v/v) heat-inactivated fetal bovine serum (FBS; HyClone, Oud-Beijerland, The Netherlands), at 37°C with 5% (v/v) CO2 in a humidified atmosphere. No antibiotics were used. Cells were split 1:6 twice weekly and fed 24 h before each experiment.
Differentiation into myotubes was achieved by culturing preconfluent cells (85% confluence) in medium containing 2% FBS for 6–8 d. The reduced serum level allowed for cell-to-cell fusion and the formation of myotubes expressing myogenin, a marker of muscle differentiation (6). Treatments were always started at day 6 from medium switching.
In this study, myoblasts and myotubes were treated with either vehicle (ethanol, EtOH) (0.01%–2% v/v) or salmeterol (0.1–20 μM) for different times defined as short term (6 h), midterm (12–72 h), and long term (>3 d). In mid- and long-term treatments, salmeterol was replaced every 48 h. Preliminary experiments verified that, at the final dilution used in the culture medium, the ethanol did not have any specific effect compared to untreated cells (data not shown). The β2 agonist was freshly prepared in EtOH vehicle (Sigma-Aldrich, St. Louis, MO) at a stock concentration of 1 mM and was further diluted in the culture medium. Because salmeterol therapeutic dose produces plasma drug concentrations at the nanomolar level (13), and the concentration of 2–3 μM in biological samples is referred as doping abuse of the drug (40), all concentrations used in this study were defined as “high” and/or “supratherapeutic” concentrations.
All chemical reagents, unless specified otherwise, were purchased from Sigma-Aldrich.
Cell Viability and Cell Growth Assessment by MTS Assay
Cell survival was measured using a 3-(4,5-dimethylthiazol-1)-5-(3-carboxymeth-oxyphenyl)-2H-tetrazolium (MTS) assay (Promega, Madison, WI). The assay is based on the enzymatic cleavage and conversion of the soluble yellow dye MTS to the water-insoluble purple formazan in living cells. Using this test, we can distinguish between two separate response components in the bioassay. One accounts for the relatively high background of the assay and is probably due to formazan produced by reducing substances (e.g., glutathione) in the assay medium. The other response component, which is dependent on the presence of an intermediate electron acceptor, phenazine methosulfate, reflects the metabolic activation of the cells. Therefore, the amount of formazan produced is proportional to the number of living cells and their relative metabolic rate. Cells were cultured in 96-well plates at a density of 1 × 104 cells per square centimeter and treated with salmeterol (0.1–10 μM for 6, 12, 24, and 48 h). Then, 20 μL of MTS was added to each well, and the plates were incubated at 37°C for 2 h. The absorbance of the converted dye was measured at 490 nm using a plate reader (680 Microplate Reader; Bio-Rad Laboratories, Inc., Hercules, CA) and is expressed as the optical density (OD) value. Four independent experiments were performed.
Assessment of Cell Viability Using Trypan Blue Exclusion Assay
The susceptibility of cultured cells to the cytotoxic effects of salmeterol was evaluated using the Trypan blue exclusion assay (Sigma-Aldrich). The cells that exclude the dye are viable because the chromophore is negatively charged and does not enter the cell unless the membrane is damaged. Myoblasts cultured in Dulbecco modified Eagle medium with 10% FBS were plated onto six-well plates at a density of 1 × 104 cells per square centimeter the day before stimulation. The cells were then incubated with the indicated concentration of salmeterol (0.1–10 μM) for different times before collection and analysis. Cells already detached were collected with the culture media and added to those subsequently obtained by trypsinization. Once centrifuged (1100g for 10 min) and resuspended in 1 mL of phosphate-buffered saline (PBS), 50 μL of cells’ suspension was treated with an equal volume of reagent (trypan blue solution, 0.4%; Sigma-Aldrich) and immediately counted by a hemocytometer (Bürker counting chamber; Brand GmbH, Wertheim, Germany). The number of nonviable cells (stained cells) was counted using a microscope and is expressed as the percentage of the total number of counted cells. Four independent experiments were performed in triplicate.
In experiments on long-term exposure to salmeterol, the cells were plated onto 25-cm2 flasks at a density of 3.2 × 103 cells per square centimeter and maintained in a proliferative state by trypsinization and subculturing at days 3, 6, 10, 13, and 17 from the beginning of the culture. At each experimental point, the total number and the percentage of living myoblasts were evaluated (trypan blue assay) before reseeding the cells at the same density onto T25-cm2 flasks in culture media containing salmeterol (2.5 or 5 μM) or vehicle. This procedure allows the maintenance of both culture conditions and the myoblasts’ proliferative state, being all cultures harvested within the 60% to 80% of confluence for the entire duration of the protocol (17 d). At each subculturing, the fold increase or decrease in cell number was calculated and taken into account for the determination, at each point, of the total cell number. The data show the results from three independent experiments.
RNA Extraction and Reverse Transcription–Polymerase Chain Reaction
To examine ADRB1 and ADRB2 expression in our cellular models, reverse transcription–polymerase chain reaction (RT–PCR) was performed with RNA extracted from the cultured C2C12 and L6C5 myoblasts. Briefly, total RNA was isolated using TRIzol reagent (Invitrogen, Life Technologies) according to the manufacturer’s recommendations, dissolved in RNAse-free water, and kept at −80°C until use. To remove any contaminating genomic DNA, the RNA was treated with DNase during the RNA isolation. For the synthesis of complementary DNA, 3 μg of total RNA was reverse-transcribed for 1 h at 42°C in a reaction mixture (20 μL) containing 50 mM Tris–HCl (pH 8.3), 40 mM KCl, 3 mM MgCl2, 1 mM dithiothreitol, 0.5 mM of each dNTP, 1.25 μg of oligo(dT), and 500 U of SuperScript II (Invitrogen, Life Technologies). Controls omitting the reverse transcriptase or template were performed by the addition of nuclease-free water. The obtained complementary DNA (3 μL) was used to amplify ADRB1, ADRB2, and GAPDH using following primers: ADRB1 sense (mouse and rat) 5′-GAGCTCTGGACTTTCGGTAGA-3′ and antisense (mouse and rat) 5′-GGCACGTAGAAGGAGACGAC-3′; ADRB2 sense (mouse and rat) 5′-AAGAATAAGGCCCGAGTGGT-3′ and antisense (mouse and rat) 5′-GTCTTGAGGGCTTTGTGCTC-3′; GAPDH sense (mouse and rat) 5′-ACCACAGTCCATGCCATCAC-3′ and antisense (mouse and rat) 5′-TCCACCACCCTGTTGCTGTA-3′. The amplification protocol was as follows: 35 cycles of denaturation at 94°C for 30 s, annealing at 55°C for 30 s, and extension at 72°C for 1 min. Because the level of glyceraldehyde phosphate dehydrogenase (GAPDH) is independent of the degree of confluence of these cultures, it was used as internal control to normalize ADRB1 and ADRB2 gene expression. The amplicon size was 313 bp for ADRB1, 383 bp for ADRB2, and 450 bp for GAPDH. The PCR products were separated using agarose gel electrophoresis (1% agarose; Bio-Rad Laboratories, Inc.) to confirm the target product of PCR for each reaction. Images of ethidium bromide–stained agarose gels were acquired, and the quantification of bands was performed using the Gel-ImageJ software (W. S. Rasband, ImageJ; US National Institutes of Health, Bethesda, MD). The results are expressed as the ratio between the OD of the adrenergic β1- or β2-receptor and GAPDH. RNA extracted from HL-1 mouse cardiac cells was used as a positive control for the detection of ADRB1 and as a negative control for ADRB2 expression.
C2C12 and L6C5 cells were grown on 12-mm glass coverslips (neoLab Migge Laborbedarf-Vertriebs GmbH, Heidelberg, Germany) in culture dishes. After treatment with different salmeterol concentrations (5–10 μM) for 48 or 72 h, cells from each group were washed with PBS and fixed with 4% formaldehyde at 4°C for 1 h. The fixed cells were then stained with 10 μg·mL−1 Hoechst 33258 (Invitrogen, Life Technologies) for 15 min in the dark to stain nuclei. The number of mitotic figures was counted using a fluorescence microscope (Eclipse E600; Nikon Instruments, Melville, NY). For each treatment, the mitotic index (mitotic figures/total cells) was calculated in three replicate wells. Three independent experiments were performed.
A total of 3.2 × 103 cells per square centimeter were plated in 10-cm plates (protein content) or T75 (enzymatic activity) flasks and treated as reported above. After each treatment, the cells were trypsinized, counted, and centrifuged at 1100g for 10 min at room temperature. The cells were then lysed in extraction buffer (50 mM Tris–acetate, 250 mM sucrose, pH 7.5) supplemented with 1 mM phenylmethylsulfonyl fluoride (PMSF) and protease inhibitor cocktail (P8340; Sigma-Aldrich). The cells were then gently sonicated for a total of 20 s (10 s twice) on ice using a Vibra-Cell CV 18 SONICS VX 11 (Sonics & Materials, Newtown, CT), and the resulting lysates were centrifuged at 14,000 rpm for 10 min at 4°C. The supernatants were tested for protein content using the Bradford method (Sigma-Aldrich) and then analyzed spectrophotometrically (Perkin Elmer Lambda 25, Fremont, CA) for enzymatic activities under substrate saturating condition at experimentally determined dilution factors following published protocols (10).
GAPDH activity measurements were performed by quantifying the reduction of NAD+ (detected at 340 nm) at 30°C in an assay mixture containing 50 mM sodium phosphate (pH 8.9), 1 mM EDTA, 1 mM oxidized NAD+, and 1 mM glyceraldehyde-3-phosphate with a 100-μL sample (minimum 100 μg of protein). Millimolar extinction coefficient, [Latin Small Letter Open E]340 = 6.22.
LDH activity measurements were performed by quantifying the reduction of NAD+ (measured at 340 nm) at 30°C in an assay mixture containing 0.2 M Tris–HCl (pH 7.6), 7 mM oxidized NAD+, and 55 mM lactate with a 20-μL sample (minimum 30 μg protein). Millimolar extinction coefficient, [Latin Small Letter Open E]340 = 6.22.
Citrate synthase (CS) activity measurements were measured using the Ellman reagent (5,5′-dithiobis-(2-nitrobenzoic acid); DTNB) at 412 nm in an assay mixture containing 1 mM DTNB in 1 M Tris–HCl (pH 8.0), 10 mM acetyl-CoA, 10 mM oxalacetate with a 50-μL sample. Millimolar extinction coefficient, [Latin Small Letter Open E]412 = 13.6.
3-OH Acyl-CoA Dehydrogenase
3-OH acyl-CoA dehydrogenase (HAD) activity measurements were performed by quantifying the oxidation of NADH (detected at 340 nm) at 30°C in an assay mixture containing 50 mM triethanolamine–HCl (pH 7.0), 4 mM EDTA, 0.04 mM NADH, and 0.015 mM S-acetoacetyl-CoA with a 50-μL sample (minimum, 100 μg protein). Millimolar extinction coefficient, [Latin Small Letter Open E]340 = 6.22.
Alanine transglutaminase (ALT) activity measurements were performed by quantifying the oxidation of NADH (detected at 340 nm) at 30°C in an assay mixture containing 0.51 M L-alanine, 136 U·mL−1 LDH, 14.5 mM α-ketoglutaric acid, and 2.2 mM NADH in 80 mM Tris–HCl (pH 7.8) with a 50-μL sample. Millimolar extinction coefficient, [Latin Small Letter Open E]340 = 6.22.
One unit of enzymatic activity was defined as the amount of enzyme that forms 1 μmol of product per minute per milligram of protein tested.
Cell Apoptosis Assay
C2C12 and L6C5 cells were grown on 12-mm glass coverslips (neoLab Migge Laborbedarf-Vertriebs GmbH) in culture dishes. After treating with different salmeterol concentration (5–10 μM) for 48 or 72 h, cells from each group were washed with PBS and fixed with 4% formaldehyde at 4°C for 1 h. The fixed cells were then stained with 10 μg·mL−1 Hoechst 33258 for 15 min in the dark to stain nuclei. The cells were observed and photographed using a fluorescence microscope (Eclipse E600; Nikon Instruments). Apoptotic cells were identified as those with a nucleus containing brightly stained condensed chromatin or nuclear fragmentation, compared with normal blue nuclei with an organized structure. For each experimental condition, four separate cell populations were prepared. At least 200 cells in five randomly selected fields were counted and quantified for each cell population and each experimental condition (total 1000 cells per population). The apoptotic index (the percentage of apoptotic cells) was calculated as the number of apoptotic cells divided by total cells counted × 100.
The terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay was used to detect apoptotic DNA breaks in situ using the Detection Kit (Roche Applied Sciences, Penzberg, Germany) following the manufacturer’s instructions. Cells attached to glass slides in culture dishes were analyzed using the TUNEL assay and an Olympus BX41 fluorescence microscope (Olympus Microscopy, Southend-on-Sea, UK). This assay preferentially labels DNA strand breaks that are generated during apoptosis. Because the presence of nonfused apoptotic myoblasts determined by the differentiation process itself makes difficult to perform the apoptotic analysis at asingle cell level, the Hoechst and TUNEL analysis was not performed in C2C12 and L6C5 myotubes.
Images were acquired using a digital camera (Infinity 1, K100; Lumenera, Ottawa, Ontario, Canada) and analyzed using the Image Analysis System (Alexa Soft s.a.s., Florence, Italy).
Preparation of Cell Homogenates and Western Blot Analysis of Apoptotic Markers
After the indicated treatment, cells were lysed, and total cellular proteins were extracted by scraping into ice-cold lysis buffer (20 mM Tris pH 7.5, 150 mM NaCl, 2 mM EDTA, 1 mM sodium orthovanadate, 100 μM PMSF, 10 μg·mL−1 leupeptin, 10 μg·mL−1 aprotinin, 5 μg·mL−1 pepstatin, 50 mM NaF, 1 nM okadaic acid, 1% Triton X-100, and 10% glycerol). The protein content of the extract was determined using the Bradford assay (Sigma-Aldrich). The extracted proteins extracted were used immediately or divided and stored at −80°C until used.
Cellular proteins (10–20 μg in 25 mM Tris–HCl pH 8, 0.5% SDS, 0.05% 2-mercaptoethanol, 2.5% glycerol, and 0.001% bromophenol blue) were denatured at 100°C for 5 min, subjected to SDS–PAGE in a 8%–12% polyacrylamide gel, and then electroblotted onto a polyvinylidene fluoride membrane at 130 V for 1 h. The blots were blocked with 5% nonfat dry milk (Bio-Rad Laboratories, Inc.) and then incubated with anti–caspase-8, anti–caspase-9, anti–Smac/Diablo, anti–Bcl-xL, or anti–poly (ADP-ribose) polymerase (PARP) antibodies (Santa Cruz Biotechnology, Santa Cruz, CA). After washing and incubating with a horseradish peroxidase–conjugated secondary antibody, the oxidation of luminol induced by the peroxidase was detected by the enhanced chemiluminescence method (Amersham Biosciences, GE Healthcare Europe GmbH, Glattbrugg, Switzerland). Bands were quantified using ImageJ software (US National Institutes of Health; http://rsb.info.nih.gov/ij, 1997–2008). The expression of β-actin (Sigma-Aldrich) or COXIV (Santa Cruz Biotechnology) was used to normalize the data.
Cells were removed by trypsinization in 0.25% trypsin and harvested by centrifugation at 1000g for 10 min. Floating cells were also collected in the same tubes. The cell pellet was washed twice with ice-cold PBS and centrifuged. The pellet was then suspended in five volumes of buffer A [20 mM HEPES–KOH (pH 7.5), 1.5 mM MgCl2, 10 mM KCl, 1 mM dithiothreitol, 1 mM Na-EDTA, 1 mM Na-EGTA, and 0.1 mM PMSF containing 250 mM sucrose] with protease inhibitors [5 μg·mL−1 pepstatin A, 10 μg·mL−1 leupeptin, 2 μg·mL−1 aprotinin, and 25 μg·mL−1 N-acetyl-leu-leu-norleucine]. After incubating on ice for 15 min, the cell suspension was gently homogenized using a glass Dounce homogenizer (20–30 strokes), and the cell lysates were checked by trypan blue staining. After centrifuging twice at 750g for 10 min at 4°C, the supernatant was collected and centrifuged at 14,000g for 15 min at 4°C. Next, the supernatant was collected, and the resulting mitochondrial pellets were resuspended in buffer A. The cytoplasmic and mitochondrial fractions were assessed for protein content using the Bradford method (Sigma-Aldrich). The purity of both cellular fractions was preliminarily ascertained by Western blot analysis performed using COXIV as a mitochondrial marker and β-actin as a cytosolic marker (data not shown).
For indirect immunofluorescence, the cells were grown on 30-mm glass coverslips (neoLab Migge Laborbedarf-Vertriebs GmbH) in six-well plates (1 × 104 cells per square centimeter) or on 15-mm glass coverslips (neoLab Migge Laborbedarf-Vertriebs GmbH) in 12-well plates (1 × 104 cells per square centimeter). C2C12 and L6C5 myoblasts were treated with 2.5, 5, or 10 μM salmeterol for 48 h. After the experimental treatments, the cells were washed with PBS and fixed in 4% paraformaldehyde diluted 1:1 with PBS/0.3% Triton X-100 for 20 min. The cells were washed and then blocked in 5% normal goat serum/1× PBS/0.3% Triton X-100 for 1 h at room temperature.
The cells were incubated overnight at 4°C with a rabbit anti–Smac/DIABLO antibody (1:200; Santa Cruz Biotechnology). After incubation, the cells were rinsed three times with PBS for 10 min each and then incubated with a fluorescein isothiocyanate–conjugated goat antirabbit antibody (1:500; Santa Cruz Biotechnology) for 1.5 h at room temperature. After this incubation, the cells were rinsed three times with PBS for 5 min each. A control stained with only the secondary antibody was performed to exclude background fluorescence. The stained cells were then stained with 10 μg·mL−1 Hoechst 33258 for 15 min in the dark to stain nuclei. Images were obtained using an Eclipse E600 (Nikon Instruments) fluorescence microscope and a digital camera (Infinity 1, K100; Lumenera) and analyzed using the Image Analysis System (Alexa Soft s.a.s.).
Quantitative RT–PCR was used to evaluate the salmeterol-mediated modulation of ADRβ1 and ADRβ2 expression. The purity, integrity, and yield of RNA were monitored using microcapillary electrophoresis (Bioanalyzer 2100; Agilent Technologies, Foster City, CA) using the RNA 6000 LabChip kit. RNA (1 μL) was treated with Amplification Grade DNAse I (Invitrogen, Life Technologies) and reverse-transcribed using the SuperScript III Kit (Invitrogen, Life Technologies). Quantitative PCR was performed using an ABI PRISM 7000 Light Cycler (Applied Biosystems, Life Technology Corporation, Carlsbad, CA) using the SYBR Green qPCR SuperMix for ABI Prism (Invitrogen, Life Technologies) as indicated by the manufacturer’s instructions. All primers were optimized for real-time amplification by verifying the production of a single amplicon in a melting curve assay and the efficiency in a standard curve amplification (>98% for each pair of primers). Messenger RNA (mRNA) expression levels were normalized to the 18S ribosomal RNA control gene after determining that there was no change in 18S RNA in response to the treatments. The relative level for each gene was calculated using the 2−ΔΔCt method and is reported in arbitrary units. In all experiments, each sample was analyzed in triplicate. The sequences of the primers are as follows:
rat ADRB1 (GenBank NM012701) sense 5′→3′ CCGCTGGGAGTACGGCTCCT and antisense 5′→3′ CCCGCCACCAGTGCATGAGG;
rat ADRB2 (GenBank NM012492) sense 5′→3′ AGCAGGATGGGAGGAGCGGG and antisense 5′→3′ GGTTGGCCCGGATGACGTGG;
rat 18S (GenBank X01117) sense 5′→3′ CGCGGTTCTATTTTGTTGGT and antisense 5′→3′ AGTCGGCATCGTTTATGGTC;
mouse ADRB1 (GenBank NM007419) sense 5′→3′ GTAACGTGCTGGTGATCGTG and antisense 5′→3′ AAGTCCAGAGCTCGCAGAAG;
mouse ADRB2 (GenBank NM007420) sense 5′→3′ GAGCACAAAGCCCTCAAGAC and antisense 5′→3′ GTTGACGTAGCCCAACCAGT;
mouse 18S (GenBank X00686) sense 5′→3′ CCCTGCCCTTTGTACACACC and antisense 5′→3′ CGATCCGAGGGCCTCACTA.
All values are expressed as mean ± SDs. Data normality were determined using the Kolmogorov–Smirnov test and statistical differences were assessed using an independent Student’s t-test for two-group comparison or dependent t-test for comparison between the same group at different time points. For data involving three or more groups and for measuring the effect of two parameters, such as treatment and time point, data were subjected to two-way ANOVA corrected for multiple comparisons with Bonferroni post hoc assessment. Analysis were performed using the statistics program SPSS (Version 15.0 for Windows; SPSS Inc., Chicago, IL). Data were considered statistically significant at P < 0.05.
Expression of β1- or β2-adrenergic Receptor mRNA in C2C12 and L6C5 Skeletal Muscle Cells
To determine the expression pattern of the β-adrenergic receptors, we performed semiquantitative RT–PCR on RNA isolated from C2C12 and L6C5 myoblasts. Both cell lines expressed the β2-isoform of the receptor and very low levels of the β1-isoform (Fig. 1A). HL-1 cardiomyocyte extracts were used as a negative control for β2 expression and a positive control for β1 expression.
Salmeterol Treatment Impairs the Growth and Viability of Proliferating C2C12 and L6C5 Myoblasts
To evaluate the effect of salmeterol on cell growth and/or viability, we first analyzed the time-dependent response of proliferating myoblasts to different salmeterol concentrations (0.1–10 μM) using the MTS metabolic assay. We observed different dose- and time-dependent effects on skeletal muscle cells. After 6 h of treatment, there was a dose-dependent increase in the absorbance in both cell lines (P < 0.05 compared control cells for doses greater than 1μM), and this effect was maintained up to 24 h of exposure. After 24 h, cells treated with the highest concentration of salmeterol showed a dramatic decrease in the OD values, indicating a significantly lower number of viable cells compared with the control cultures (Figs. 1B, C; P < 0.01). The kinetics of the impairment in cell viability were the same for the two cell lines.
Treatment with 10 μM salmeterol for 48 h resulted in significant toxicity and a 40% and 46% reduction in cell viability for C2C12 and L6C5 myoblasts, respectively, compared with the control group. Because the doubling time of these cells is between 19 (C2C12) and 22 h (L6C5), we performed the trypan blue exclusion assay to determine the extent to which the variation in the OD values is the result of variations in cell number. As shown in Figures 1D and E, after 6 h of treatment with 5 μM (C2C12 cells) or 10 μM (L6C5 cells) salmeterol, no changes in the number of viable cells were found. In contrast, in C2C12 myoblasts, treatment with either 5 or 10 μM salmeterol for 24 h was able to significantly reduce the number of viable cells (35.6% and 39.3%, respectively; P < 0.05) and increase cell death (3.2 and 3.5 times) compared with the control cells. In L6C5 cells, while 5 μM salmeterol did not significantly affect the number of viable cells, treatment with 10 μM salmeterol induced a statistically significant (P < 0.01) increase in cell death (4.9 times over the control) and a decrease in the number of viable cells (60% compared with the control cells; P < 0.01).
To assess whether long-term treatment with lower concentrations would also exert cytotoxic effects on C2C12 and L6C5 myoblasts, we evaluated the effects of exposure to 2.5–5 μM salmeterol for 3–17 d. The observed number of viable cells detected using the trypan blue exclusion assay are reported in Figure 2. In both cell lines, salmeterol caused a dose-dependent inhibition of cell growth. L6C5 myoblasts seemed more sensitive to long-term exposure, showing cell growth arrest after 3 d with 2.5 and 5 μM salmeterol (P < 0.01; Fig. 2B), while after 3 d in 5 μM salmeterol, the C2C12 myoblasts showed a complete inhibition of cell growth (P < 0.05). The decrease in cell number seemed to be related to the inhibition of cell growth rather than to increased cell death. While we observed a dose- and time-dependent increase in the proportion of trypan blue–positive cells in both cell lines, the values were not statistically significant compared with untreated cultures (data not shown).
These data clearly demonstrate that, in myogenic cells, (a) short-term exposure to different concentrations of salmeterol rapidly increases the cellular metabolic rate without increasing cell number, (b) long-term exposure to relatively low concentrations of salmeterol (2.5–5 μM) results in strong cytostatic effects, and (c) midterm exposure (48 h) to high salmeterol concentrations (10 μM) has both cytostatic and cytotoxic effects.
Short-term Treatment with Salmeterol in C2C12 and L6C5 Modulates the Activity of Metabolic Enzymes
Because the MTS assay indicates an increase in the metabolic rate after short-term exposure to salmeterol, we determined whether salmeterol treatment could modulate the activity of key enzymes involved in energy metabolism. The activity of several enzymes, such as GAPDH, LDH, CS, HAD, and ALT, was analyzed either in proliferating or differentiated C2C12 and L6C5 skeletal cells treated with 10 μM salmeterol for 6 h.
Short-term treatment maximized the metabolic oxidative pathway in myoblasts from both cell lines; we found an increase in the activity of GAPDH (62% for C2C12 and 19% for L6C5), CS (aerobic metabolism; 23% for C2C12 and 15% for L6C5), HAD (aerobic lipidic metabolism; 57% for C2C12 and 78% for L6C5), and ALT (amino acid metabolism; 40% for C2C12) compared with control cells. In addition, the LDH activity decreased in both cell lines with a greater effect in L6C5 myoblasts (15% decrease in C2C12 cells and 63% decrease in L6C5 cells; Figs. 3A, C). C2C12 myotubes showed the same response as myoblasts, with a general increase in aerobic metabolism (a 24%, 54%, 37%, and 10% increase in GAPDH, CS, HAD, and ALT activity, respectively, and an 11% decrease in LDH activity). In contrast, L6C5 myotubes demonstrated a pronounced activation of the anaerobic pathway, with an increase in the activity of GAPDH (56%) and LDH (anaerobic metabolism; 34%), a decrease in the activity of CS (aerobic metabolism; 16%) and HAD (aerobic lipidic metabolism; 5%), and no change in the activity of ALT (amino acid metabolism) compared with control cells (Figs. 3B, C). We asked then if the metabolic changes induced by salmeterol in C2C12 and L6C5 myoblasts and myotubes, beside the differences in the enzymatic activities, could result in a common cellular response. Indeed, under the same experimental condition used for the metabolic analysis, both myoblasts and myotubes showed a significant increase (P < 0.05) in total protein content (Bradford assay) (C2C12 myoblasts: control 125 ± 8.7 μg vs salmeterol 157 ± 14.3 μg; L6C5 myoblasts: control 101 ± 8.4 μg vs salmeterol 124 ± 9.5 μg; C2C12 myotubes: control 15.0 ± 0.94 mg vs salmeterol 17.5 ± 0.90 mg; L6C5 myotubes: control 11.9 ± 0.73 mg vs salmeterol 14.9 ± 0.69 mg), suggesting a possible anabolic effect of the drug.
Midterm Exposure to High Salmeterol Concentrations Induces Mitotic Arrest in Myoblasts and Cellular Apoptosis in Both Myoblasts and Myotubes
Preliminary studies have shown that β-AR agonist use may be associated with the induction of apoptosis (3). As cited above, the analysis of cell growth and viability revealed a decrease in the cell number after exposure to salmeterol. This finding suggests that cell growth arrest and/or cell death may occur because of the treatment. Therefore, we evaluated whether salmeterol inhibited mitosis or induced apoptosis in myogenic cells. We performed two specific morphological assays, Hoechst staining and TUNEL, which are able to distinguish mitotic and/or apoptotic cells. As shown in Figures 4A and B, salmeterol treatment (5 and 10 μM) for 48 and 72 h completely blocked mitosis in both C2C12 and L6C5 myoblasts in a dose-dependent manner (P < 0.05). Moreover, compared with control cells that demonstrated a normal nuclear morphology, the treated cells showed morphological changes typical of apoptosis, such as fragmented or pyknotic nuclei. After treatment with 10 μM salmeterol, 60% of C2C12 cells (Fig. 4C) and 76% of L6C5 (Fig. 4D) cells seemed to be undergoing apoptosis, a statistically significant difference compared with untreated cultures (6% and 7%–8% apoptosis, respectively; P < 0.05). These data were further confirmed in cells treated for 48 h with 2.5, 5, or 10 μM salmeterol using the TUNEL assay. There was a dose-dependent increase in TUNEL-positive nuclei, with 25% of C2C12 cells (Fig. 4E) and 29% of L6C5 cells (Fig. 4F) TUNEL-positive after treatment with 10 μM salmeterol compared with 3% and 4% of untreated cells, respectively. The difference in the percentages of apoptotic cells determined using Hoechst staining and the TUNEL assays is likely due to the specific properties of these assays. While the Hoechst staining is very sensitive to the DNA conformation and chromatin state in cells, and thus detects gradations of nuclear damage in a nonspecific manner, the TUNEL reaction preferentially labels DNA strand breaks that are generated during apoptosis. This allows for the discrimination of apoptosis from necrosis (12).
Next, we investigated the molecular mechanism by which salmeterol induced apoptosis in proliferating and differentiated cells. The apoptotic program involves two major pathways. The extrinsic pathway relies on the action of cell surface death receptors and caspase-8, whereas the intrinsic pathway is mediated by caspase-9 through the mitochondrial death cascade and the activation of many other proteins such as PARP, the Bcl-2 family of proteins, and Smac/DIABLO (17,41). Given that these proteins play an important role in apoptosis, we determined the effect of treatment with 2.5, 5, or 10 μM salmeterol for 48 h on their expression in both proliferating and differentiated C2C12 and L6C5 cells treated. While the induction of pro–caspase-8 cleavage seemed to be unaffected in both myoblasts and myotubes, we observed a dose-dependent increase in caspase-9 activation in C2C12 and L6C5 skeletal myoblasts and myotubes (Fig. 5A). Similar to previously reported results with other apoptotic stimuli (6), differentiated C2C12 and L6C5 myotubes seemed more resistant to salmeterol-induced apoptosis than myoblasts. In proliferating cells, the increased pro–caspase-9 cleavage was detectable after treatment with 2.5 μM salmeterol, while in myotubes, the cleaved fragment was clearly evident after treatment with 10 μM salmeterol. Moreover, exposure of C2C12 myotubes to 10–20 μM salmeterol and L6C5 myotubes to 5–10 μM salmeterol for 48 h induced the dose-dependent activation of PARP, and this was paralleled by the decreased expression of the antiapoptotic protein Bcl-xL (Fig. 5B).
During apoptosis, mitochondria also undergo changes in structure and localization. The alterations frequently observed are pyknosis, a reduction in size, and mitochondrial grouping in the perinuclear zone. Smac/DIABLO is normally localized in the mitochondria, so we analyzed its subcellular localization using immunofluorescence. In untreated C2C12 and L6C5 myoblasts, Smac/DIABLO expression had a normal “spider web” distribution throughout the cell, distant from the nucleus. In contrast, after salmeterol treatment, we observed the cytoplasmic diffusion of Smac/DIABLO and a clustered mitochondrial distribution around the nucleus (Fig. 6A). This observation is quantified in the graphs presented in Figure 6B, which shows the percentage of cells where Smac/DIABLO was diffusely expressed in the cytoplasm as a result of the activation of the apoptosis (C2C12: control 1.3% ± 0.4%, 10 μM 20% ± 3%; L6C5: control 2% ± 0.7%, 10 μM 18% ± 2%; P < 0.05). The results from indirect immunofluorescence assay were confirmed using Western blot analysis on subcellular fractions, which confirmed that, after treatment with 10 μM salmeterol, there was a significant (P < 0.05) decrease in mitochondrial Smac/DIABLO and an increase in cytosolic Smac/DIABLO in both C2C12 (Fig. 6C) and L6C5 (Fig. 6D) myoblasts. Therefore, these data suggest that high doses of salmeterol induce apoptosis in our skeletal muscle models via a mitochondrion-mediated pathway.
Salmeterol Does Not Modulate the Expression of β1- or β2-adrenergic Receptor mRNA
To analyze the β-adrenergic receptors modulation after treatment with salmeterol, we performed quantitative RT–PCR. In contrast to the β2-adrenergic receptor desensitization that has been observed in several tissues after long-term treatment with β2 agonists, neither C2C12 nor L6C5 myoblasts treated with salmeterol demonstrated decreased expression of the β2-receptor. Indeed, quantitative analysis showed that treatment with 2.5 μM salmeterol for different times (6–72 h) did not cause a significant increase in ADRB2 expression in C2C12 myoblasts (see Figure, Supplemental Digital Content 1A, http://links.lww.com/MSS/A96, ADRB2 expression by quantitative RT–PCR). In L6C5 cells (see Figure, Supplemental Digital Content 1B, http://links.lww.com/MSS/A96, ADRB2 expression by quantitative RT–PCR), ADRB2 expression also seemed to be unaffected. In addition, exposure to salmeterol did not affect the expression of ADRB1, which remained undetectable at the mRNA level (data not shown).
β2 agonists are often used to treat asthma and COPD owing to their ability to relax pulmonary smooth muscle; however, there is evidence that therapeutic use can have adverse effects, some of which may be severe (27,32). It is known that the in vivo administration of β-AR agonists leads to the activation of caspase-3 and increases the sensitivity of myocyte DNA to denaturation and strand breaks, both of which is a hallmark of apoptosis (4). The illicit use of these drugs could cause the cytotoxic effects mentioned above, especially when they are administered in doses that exceed the normal therapeutic ones. Moreover, the increased number of athletes using β2-adrenergic receptor agonists (at higher percentages than asthmatic individuals), in addition to the lack of information regarding the effects of abuse of these compounds, has prompted the World Anti-Doping Agency to restrict β2 agonist use, except for specific medical reasons (40).
The main aim of this work was to determine the effect of different doses of salmeterol on cell growth, metabolism, and activation of apoptotic pathways using in vitro skeletal muscle models, the C2C12 mouse and L6C5 rat muscle cell lines, either under proliferating or differentiated states. Furthermore, to verify the effectiveness of these models, we also evaluated the expression of the two most important proteins involved in the β-agonist response, the β1- and β2-receptors, using the HL-1 mouse myocardial cell line as a control. Our data are in agreement with other previous results and demonstrated that the expression of the β1-receptor is increased in the cardiac tissue, accounting for approximately 60%–80% of the β-adrenergic receptor population. On the other hand, in striated muscle cells, the β2-receptor is the most highly expressed, accounting for 80%–90% of the β-adrenoreceptors (2).
The analysis of skeletal myoblast cell growth after treatment with salmeterol (6–48 h) in the MTS metabolic assay showed a clear and significant increase in absorbance, starting after 6 h of exposure to as little as 1 μM salmeterol in both cell lines. The length of the cell cycle depends on the organism and cell type (7). Although immortalized cell lines can have a shorter cell cycle compared with primary cells, the doubling times of C2C12 and L6C5 myoblasts are not consistent with an increase in cell number during a 6-h treatment.
Because this assay is based on the conversion of the MTS tetrazolium compound into a colored formazan product that is presumably accomplished by NADPH or NADH produced by dehydrogenase enzymes in metabolically active cells (1), the results found suggested an enhanced metabolic rate rather than an effect on the growth rate of the cells.
This hypothesis was further confirmed using the trypan blue exclusion assay. After 6 h of salmeterol treatments, the number of cells was similar in all cultures, but the metabolism of the cells was altered.
For this reason, we focused on the short-term effect of salmeterol on myoblast’s metabolism with respect to prolonged exposure, in which MTS signal started to decline.
β-AR agonist administration has been found to modulate oxidative metabolism, energy expenditure, lipolysis (16), glucose transport (28), glucose oxidation (19), and mitochondrial morphology (38) in skeletal muscle.
In the current study, we found that salmeterol administration significantly modified the activity of enzymes involved in cell metabolism. Our data suggest an increase in oxidative metabolism in both myoblast cell lines and in C2C12 myotubes, as illustrated by the increased CS activity paralleled with the decreased LDH activity. Conversely, L6C5 myotubes showed a significant increase in anaerobic metabolism, but this trend can be explained by differences in the principal metabolic pathways active in these cells after differentiation (37). As demonstrated with other β-agonist (18), our results show that short-term salmeterol treatment is able to increase the total protein content in both myoblasts and myotubes culture. The data may suggest an anabolic effect of salmeterol on skeletal muscle. This important issue actually overcomes the aims of the current research and deserves further dedicated studies to clarify the molecular pathway/s possibly involved.
Despite short-term treatment, it seems able to increase skeletal muscle metabolism; on the other hand, for mild- and long-term treatments, salmeterol was increasingly toxic in a dose- and time-dependent manner. In agreement with the work by Stewart et al. (36) on human airway smooth muscle cell after β-agonists prolonged treatment, our results showed that, in prolonged treatments (48–72 h), salmeterol decreased the number of cells, decreased (or completely inhibited at 10 μM) the mitotic index and increased the number of apoptotic cells in both cell lines. Moreover, long-term exposure (3–17 d) at 2.5 μM completely abolished cell growth even without cell death.
As demonstrated for clenbuterol (3), in addition to the cytostatic effect, high concentrations of salmeterol for prolonged exposures also induced apoptosis in proliferating and differentiated skeletal muscle cells. Indeed, this study analyzed for the first time the activation of the apoptotic pathway at the molecular level in skeletal muscle cells treated with a selective β-AR agonist. The increase of apoptotic cell in both cell lines found after prolonged treatments was associated with the activation of caspase-9 and PARP, the modulation of Bcl-xL, and the release of Smac/DIABLO from the mitochondrial compartment, suggesting the activation of the intrinsic apoptotic pathway.
Moreover, we performed an ADRB2 quantitative RT–PCR analyses to evaluate if the effects found could be due to a possible down-regulation of the β2-receptor induced by long-term salmeterol treatment. As expected, the mRNA levels seemed to be unaffected by drug treatment. These data are consistent with the hypothesis that the long-acting characteristics of salmeterol might be due, at least in part, to the ability of this agonist to maintain a population of functional β2-receptors through persistent increases in gene transcription, despite a prolonged low-level exposure to the agonist (22).
These different facets of salmeterol for short or prolonged treatments can be explained with a mitochondrial apoptosis induced by drastic metabolite consumption. In skeletal muscle, the increased flux of energy fuel substrates is associated with lipid accumulation, generation of reactive oxygen and nitrogen species, and cellular dysfunction leading to a glucolipotoxicity. Mitochondria are the greatest cellular sources of reactive oxygen and nitrogen species (20). In skeletal muscle, mitochondria display proton leak and reverse electron transfer in a proportion strongly dependent on mitochondrial delta-pH that is directly dependent on ATP content. In fact, complexes associated with electron transfer chain are directly linked with the tricarboxylic acid cycle activity, fatty acid oxidation, and glycolysis (25) with superoxide production from reverse electron transfer. The increase in ATP demand makes skeletal muscle more sensitive to electron pressure in the chain and to respiratory chain deficiencies, leading to superoxide production (25), which are able to damage mitochondrial DNA, diminish ATP production, and ultimately led to apoptosis.
In this context, the increase of oxidative metabolism found after salmeterol treatment in myogenic cells could induce an increase in energy substrate consumption that, inducing mitochondrial uncoupling, could lead to the cytostatic and cytotoxic effects observed.
In addition, the metabolic stimulus induced by salmeterol in skeletal muscle cells could determine an acute nutrient consumption/deprivation leading to a self-preservation states. In this situation, processes such as autophagy and/or acute atrophy through modulation of the energy sensor AMP-activated protein kinase–specific factors, such as FoxO and stress-activated protein kinase, can take place (15), reaching the threshold of apoptosis activation in cases of prolonged starvation (14,31).
Further studies will be required to clarify the link between the metabolic and proapoptotic effects of salmeterol in skeletal muscle cells. Nevertheless, we speculated that ROS induction could represent the main biological mediator of mitochondrial dysfunction. Indeed, as already demonstrated in L6C5 cells, the exposure to increased ROS concentration can determine either the cell cycle arrest, due to DNA damage, or can trigger the activation of apoptotic processes, depending on the exposure time and concentration (5). Therefore, an increase in ROS production could explain, at least partly, the cytostatic effect of midterm exposure to high salmeterol concentrations or long-term exposure to low drug concentrations.
Altogether, these data demonstrate that, although salmeterol is one of the most effective bronchodilator drugs for the treatment of asthma and has an unequivocal beneficial effect if it is used at therapeutic concentrations, exposure to supratherapeutic concentrations might result in toxicity to skeletal myoblasts and myotubes. This evidence suggests that illicit use of β2 agonists to exploit the muscle hypertrophic effect or its bronchodilator action can lead to long-term loss of muscle fibers, compromising the tissue and leading to decreased athletic performance. Indeed, as previously demonstrated in other tissues (33,42), the cytostatic effect induced by β2 agonists in proliferating cells at lower doses could produce harmful effects on skeletal muscle processes, counteracting muscle turnover and fiber remodeling. This effect could be amplified in specific muscle conditions, such as during the local acute deprivation of nutrients or after muscle trauma, both of which occur during normal athletic activity. During these events, when skeletal muscle satellite cells are normally activated to proliferate, β2 agonist use could result in an inability to support adequate fiber regeneration.
In conclusion, this study demonstrated for the first time that (a) 6 h of salmeterol treatment is effective in modulating myogenic cell metabolism, as demonstrated by an increase in the activity of enzymes involved in the oxidative pathway; (b) this effect is followed by dose-dependent cytostatic and cytotoxic effects in skeletal myoblasts; and (c) the cytotoxic effect in either myoblasts or myotubes exerted by midterm treatments with supratherapeutic concentrations of salmeterol was the result of the activation of the intrinsic apoptotic pathway, including the release of Smac/DIABLO from the mitochondria, the activation of caspase-9, the inhibition of Bcl-xL, the cleavage of PARP, and the activation of DNA fragmentation.
Guglielmo Duranti and Piergiorgio La Rosa contributed equally to the research. This work was supported by a grant from the Italian Ministry of Health (Anti-Doping Commission – grant 2006) to D.C. and S.B. and by a grant from the University of Rome “Foro Italico” (2007 program) to D.C., S.S., and P.P.
This work presents no conflicts of interest.
The results of present study do not constitute endorsement by the American College of Sports Medicine.