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Department of Applied Physiology and Kinesiology, Center for Exercise Science, University of Florida, Gainesville, Florida
Submitted 13 January 2006 ; accepted in final form 21 August 2006
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ABSTRACT |
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 TOP ABSTRACT EXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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skeletal muscle; oxidative stress; free radicals
25% of patients requiring MV (15). Clinically, this is significant because weaning accounts for >40% of the total time on the ventilator (6). The diaphragm is the principal inspiratory muscle in mammals, and evidence indicates that as few as 12 h of MV result in a decrement in diaphragmatic force production (18), and this deficit is exacerbated with increased time on the ventilator (14). MV-induced diaphragmatic contractile dysfunction is significant because respiratory muscle weakness is an important contributor to difficult weaning. Oxidative stress is a key mechanism contributing to MV-induced diaphragmatic dysfunction, and our group has detected increased levels of protein oxidation and lipid peroxidation in the diaphragm following only 6 h of MV (23, 25). Oxidative injury in the diaphragm can impair diaphragmatic contractile function in several ways. For example, oxidized proteins can be targeted by the proteasome proteolytic system, thereby accelerating diaphragmatic atrophy (5, 24). Furthermore, oxidative stress can damage muscle proteins involved in excitation-contraction coupling, thus reducing diaphragmatic force production (4, 17, 19, 21). The physiological significance of MV-induced oxidant stress in the diaphragm has been confirmed by recent experiments demonstrating that infusion of the antioxidant Trolox prevents MV-induced contractile dysfunction in the diaphragm (3).
Numerous radical-producing pathways exist in cells, and it is possible that several pathways could interact and contribute to MV-induced oxidative stress in the diaphragm. Candidate oxidant production pathways include the mitochondria, NADPH oxidase, xanthine oxidase, nitric oxide (NO) synthase (NOS), and/or an increase in reactive iron in the cell. For example, the mitochondria can produce superoxide at complexes I and III along the electron transport chain (7). Also, NADPH oxidase can catalyze the one-electron reduction of oxygen into superoxide using NADPH or NADH as the electron donor (8). Moreover, increases in calcium-activated neutral proteases (calpain) can upregulate xanthine oxidase activity, resulting in the formation of excess superoxide production (7). Moreover, skeletal muscle atrophy during prolonged periods of immobilization is associated with an increase in free, "reactive" iron (12). Free iron can react with hydrogen peroxide and superoxide to produce the highly reactive hydroxyl radical (7). Finally, NO can be produced via the enzyme NOS.
Of these potential radical-producing pathways, the present study investigates the role of the NOS pathway in MV-induced oxidative stress in the diaphragm. Three isoforms of NOS can exist in cells: 1) type I or neuronal (nNOS), 2) type II or inducible (iNOS), and 3) type III or endothelial (eNOS). All forms of NOS produce NO from L-arginine and require oxygen and NADPH as substrates, while citrulline is a by-product (20). Once produced, NO can react with superoxide to form the highly reactive peroxynitrite molecule that can promote nitration of proteins in skeletal muscle.
Regulation of NOS activity differs across the three isoforms. Specifically, the activities of the constitutive NOS isoforms (i.e., eNOS and nNOS) are calcium/calmodulin regulated, and increased intracellular free calcium levels activate both eNOS and nNOS activity (20). In contrast, iNOS is not calcium regulated and is primarily regulated at the transcriptional level in response to an inflammatory challenge (20).
Given that MV-induced oxidant stress promotes diaphragmatic contractile dysfunction, determining the pathway(s) responsible for MV-induced diaphragmatic oxidant production is important. In immobilized locomotor skeletal muscle, cytosolic levels of free calcium become elevated, and NO levels are reported to increase (13). Similarly, preliminary experiments in our laboratory suggest that prolonged MV results in increased calcium levels in the diaphragm (unpublished observations). Therefore, we formed the working hypothesis that MV promotes NO production in the diaphragm via an increase in NOS activity. Therefore, these experiments tested the hypothesis that diaphragmatic NOS levels and NO production are elevated during MV and result in the accumulation of 3-nitrotyrosine.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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Experimental protocol.
Animals were anesthetized with pentobarbital sodium (60 mg/kg body wt, injected intraperitoneally). After reaching a surgical plane of anesthesia, the acutely anesthetized control animals were killed immediately whereas the SB and MV animals received an intramuscular injection of glycopyrrolate (0.04 mg/kg body wt) and were tracheostomized utilizing aseptic techniques. Furthermore, both MV and SB animals received an intramuscular injection of glycopyrrolate (0.04 mg/kg body wt) every 2 h thereafter to reduce airway secretions. The SB animals breathed spontaneously for the 18-h duration, while the MV animals were mechanically ventilated with a volume-driven ventilator (Inspira, Harvard Apparatus, Cambridge, MA) for the same duration. In the MV animals, the tidal volume was set at 0.55 ml/100 g body wt, with a respiratory rate of 80 breaths/min and a positive end-expiratory pressure of 1 cmH2O. Appropriate adjustments in tidal volume were made (if required) within the first hour of MV using arterial blood-gas measurements as a guideline.
In the SB and MV animals, the carotid artery was cannulated to permit measurement of arterial blood pressure (BP) and the collection of blood for blood-gas analysis. Arterial blood samples were collected at hours 1, 9, and 18 of the experiment and analyzed for the partial pressures of O2 (PO2) and CO2 (PCO2) and the arterial pH using an electronic blood-gas analyzer (GEM Premier 3000; Instrumentation Laboratory, Lexington, MA). Arterial PO2 was maintained between 60 and 90 Torr throughout the experiment by gradually increasing the inspired O2 fraction using a hyperoxic gas (range 22–25% oxygen) in both SB and MV animals. Moreover, the jugular vein was cannulated for the infusion of saline and pentobarbital sodium (10 mg·kg body wt–1·h–1).
Body temperature was maintained at 37°C, and heart rate (HR) was monitored via a lead II electrocardiograph. Continuous care (hourly) during the experimental protocol included lubricating the eyes, expressing the bladder, removing airway mucus, and rotating the animal and limbs of the animal. Enteral nutrition was provided via the AIN-76 rodent diet with a nutrient composition of 15% proteins, 35% lipids, 50% carbohydrates, and vitamins and minerals (Research Diets, New Brunswick, NJ). Our planned feeding schedule was designed to provide an isocaloric diet with the nutrients administered every 2 h with a gastric tube; the total administration of 69 ml is equivalent to 69 kcal/day.
Following the experimental protocol, the diaphragm was removed, and the costal portion was dissected into multiple segments (50 mg each), quickly frozen in liquid nitrogen, and stored at –80°C for subsequent assay. Finally, to ensure that animals were free from infection during the experiment, blood samples obtained from each animal at the conclusion of the experiment were cultured to determine whether gram-positive and gram-negative bacteria were present.
NOS.
Protein levels of all three isoforms of NOS (eNOS, nNOS, iNOS) were detected by Western analysis. Crude muscle homogenate (75 µg protein) was loaded onto a 7.5% Tris-glycine SDS polyacrylamide gel and separated via electrophoresis (100 V, 1.5 h). Proteins were then transferred to nitrocellulose (2 h at 275 mA), and the membrane was blocked in 5% nonfat dry milk. The membrane was then exposed to monoclonal antibodies for eNOS, nNOS, and iNOS at a dilution of 1:500 (BD Transduction Laboratories, Lexington, KY) followed by exposure to a horseradish peroxidase-conjugated anti-mouse secondary antibody. Positive controls were included on each gel, including human endothelial cell lysate, rat cerebrum lysate, and mouse macrophage stimulated with IFN-
/LPS for eNOS, nNOS, and iNOS, respectively. Furthermore, membranes were stained with a 0.1% Ponceau stain following Western analysis to ensure equal protein loading and transfer.
Nitrate (NO3–) and nitrite (NO2–) are the stable end products of NO in vivo. Therefore, as an indicator of NOS activity, levels of total NO3– and NO2– were measured in the diaphragm with a colorimetric assay kit (Cayman Chemical, Ann Arbor, MI). Briefly, a section of costal diaphragm was homogenized 1:10 in phosphate-buffered saline (pH = 7.4). The homogenate was subjected to a series of centrifugations consisting of 10,000 g for 20 min, 100,000 g for 30 min, and 12,000 g using 10K filter tubes (Millipore, Bedford, MA) for 15 min. Next, NO3– was converted to NO2– by the addition of NO3– reductase. Finally, the addition of the Griess reagents converted the NO2– into a deep purple azo compound. Photometric measurement of the absorbance of this compound was then used to determine the concentration of NO2–.
3-Nitrotyrosine. Protein levels of 3-nitrotyrosine in the insoluble, cytosolic, mitochondrial, and membrane fractions of the cell were measured via Western analysis using the protocol described by Barreiro et al. (2). Briefly, 100 mg of diaphragm muscle were homogenized in a buffer containing 10 mM Tris-maleate, 3 mM EGTA, 275 mM sucrose, 0.1 mM DTT, and a protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO) (buffer A). One aliquot of homogenate was used for a dot blot, while the remaining homogenate was centrifuged at 1,000 g for 10 min. The pellet was then resuspended in buffer A and designated as the insoluble fraction. The supernatant was centrifuged at 12,000 g for 20 min, and the pellet was resuspended in buffer B (10 mM Tris-maleate, 0.1 mM EDTA, 135 mM KCl). The supernatant was removed, and the pellet was resuspended in buffer A and centrifuged at 12,000 g for 20 min. The pellet was resuspended in buffer A by sonication and was designated as the mitochondrial fraction. The supernatant from the last two steps was pooled and centrifuged at 100,000 g for 1 h. The supernatant was saved and designated as the cytosolic fraction, while the pellet was resuspended in buffer C (10 mM HEPES and 300 mM sucrose) and treated for 1 h in 600 mM KCl. The homogenate was then centrifuged at 100,000 g for 1 h. The pellet was resuspended in buffer A by sonication and designated as the membrane fraction. All four fractions were mixed with sample buffer and boiled for 5 min. Protein was loaded on a 4–20% Tris-glycine SDS polyacrylamide gel and separated via electrophoresis (1.5 h at 100 V). Proteins were then transferred to nitrocellulose (2 h at 275 mA) and blocked in 1% BSA. The membrane was exposed to a monoclonal antibody for 3-nitrotyrosine (Cayman Chemical, Ann Arbor, MI) followed by exposure to a horseradish peroxidase-conjugated anti-mouse secondary antibody. The presence of proteins was detected using chemiluminescence. Membranes were incubated in SYPRO ruby protein blot stain following Western analysis to control for protein loading. Levels of 3-nitrotyrosine were normalized to the associated protein band.
S-nitrosothiols. Levels of S-nitrosothiols (SNO) in the cell were measured via 4,5-diaminofluorescein (DAF-2) fluorescence detection using the protocol described by King et al. (9). Briefly, 30 mg of diaphragm muscle were homogenized in a buffer containing 250 mM sucrose, 5 mM Tris base, 50 mM KCl, 1 mM EDTA, 2 mM MgCl2, and a protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO). The homogenate was centrifuged twice at 1,000 g for 10 min, and the supernatant was collected. Protein concentration of the supernatant was determined via the Bradford protein assay. An equal volume of nondenaturing sample buffer containing 138 mM Tris·HCl, 22% glycerol, and 1% SDS was then added to the homogenate.
Gels (10%) were prerun for 30 min at 30 mA in a standard SDS buffer containing 1 mM EDTA. Following the prerun, gels were loaded and electrophoresed for 135 min at 15 mA. Gels were briefly washed in a 1 mM EDTA buffer in deionized distilled H2O. Following the wash, gels were placed in a dark chamber and incubated for 10 min with a 50 µM DAF-2 solution prepared in deionized distilled H2O. Gels were then exposed to UV light for 5 min at room temperature and imaged (Typhoon 8600, Molecular Dynamics) at an excitation wavelength of 488 nm and an emission wavelength of 530 nm. Following image acquisition, the two visible bands per sample were analyzed using Scion image software (Scion).
Measurement of diaphragmatic glutathione levels. Glutathione is the major nonprotein thiol in cells and is considered to be the most important nonenzymatic antioxidant in the cell. Since it is well established that cellular oxidative stress results in lower levels of glutathione in tissue, we measured diaphragmatic levels of GSH as a marker of oxidative stress in the diaphragm using a commercially available kit (Cayman Chemical, Ann Arbor, MI).
Statistical analysis. Comparisons between groups were made by a one-way ANOVA, and, when appropriate, a Tukey honestly significant difference test was performed post hoc. Significance was established at P < 0.05. Values are expressed as means ± SE.
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RESULTS |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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No significant differences in body weight existed between the groups before the experimental protocol, and the 18-h experimental protocol did not alter body weight in either the SB or MV groups. This indicates that our hydration and nutrition regimen was adequate. Also, note that none of the SB or MV animals tested positive for gram-positive or gram-negative bacteria, and there were no visual abnormalities of the lungs or peritoneal cavity. These results indicate that our aseptic surgical technique was successful in preventing infection.
NOS. To determine the impact of prolonged MV on diaphragmatic ability to synthesize NO, we measured both NOS protein levels and NOS activity. Our results reveal that MV did not alter diaphragmatic protein levels of eNOS or nNOS (Table 1). Furthermore, iNOS was not detected in the diaphragm of any of the experimental groups. Representative Western blots of nNOS, eNOS, and iNOS are illustrated in Fig. 1. Finally, MV did not result in increased NOS activity as measured by the levels of NO3– and NO2– in the diaphragm (Table 2).
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DISCUSSION |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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NOS does not contribute to MV-induced oxidative stress in the diaphragm. It is well established that isolated diaphragm muscle releases NO derivatives at slow rates during resting conditions and at higher rates during repetitive contractions (10, 20). Furthermore, NO production is also increased during contractile activity in locomotor skeletal muscles of rodents (1, 22).
Given that, compared with resting muscle, repetitive muscle contractile activity results in increased NO production, our prediction that prolonged MV would promote NO production in the diaphragm may appear paradoxical. Nonetheless, our hypothesis that NOS activity, NO production, and the accumulation of 3-nitrotyrosine is elevated in the diaphragm during MV was largely formulated based on a previous report indicating that immobilized skeletal muscle is associated with elevations in total calcium and knowledge that both of the constitutive isoforms of NOS (eNOS and nNOS) are calcium-activated enzymes (12). Nonetheless, our results do not support our hypothesis. Indeed, our data indicate that MV did not promote an increase in any NOS isoform or an increase in NOS activity. Furthermore, the finding that MV did not encourage an increase in the levels of 3-nitrotyrosine in the diaphragm suggests that NO production is not accelerated in the diaphragm during MV. Indeed, our analysis of 3-nitrotyrosine levels in the diaphragm was comprehensive and evaluated the nitration of proteins in a wide variety of protein pools in the cell, including the insoluble, cytosolic, mitochondrial, and membrane protein fractions. This type of comprehensive analysis is important to detect small treatment-induced changes in 3-nitrotyrosine, as shown by Barreiro et al. (2), who reported that the nitration of muscle proteins can be limited to one or two protein compartments of the muscle fiber. Therefore, the failure to separate muscle proteins into subfractions could mask increases in nitration within small pools of protein within the fiber. However, the current experiments avoided this pitfall, and, based upon our comprehensive analysis, the current results indicate that 18 h of MV are not associated with an increase in the nitration of diaphragmatic proteins. Moreover, this conclusion is supported by our findings that MV did not increase diaphragmatic levels of total NO3–/NO2– or SNO.
Our finding that MV-induced unloading of the diaphragm does not alter any of the NOS isoform levels differs from a previous report by Nguyen and Tidball (16), who reported that nNOS levels decrease in mouse locomotor skeletal muscle following 10 days of hindlimb unloading. The explanation for these divergent findings is unclear, but could be due to species differences, different durations of muscle unloading, and/or variances in the experimental model of muscle unloading. Regardless, muscle levels of NO are physiologically significant during periods of muscle inactivity, because increased production of NO in muscle fibers has been shown to decrease the activity of calpain, a calcium-activated protease in skeletal muscle. Increased calpain activity in skeletal muscle is associated with increased muscle proteolysis and may be a requirement for the degradation of muscle contractile proteins (11).
Conclusions and future directions. This is the first experiment to investigate the sources of oxidant production in the diaphragm during prolonged MV. Our results indicate that MV does not promote an increase in diaphragmatic levels of any NOS isoform, NOS activity, or cause the nitration of proteins in the diaphragm. Collectively, these results indicate that NOS-mediated production of NO is not involved in MV-induced diaphragmatic oxidant stress and contractile dysfunction. Hence, additional experiments will be required to elucidate the sources of oxidant production in the diaphragm during prolonged MV. While the current experiments rule out the possibility that NO production contributes to MV-induced oxidative stress during the first 18 h of MV, additional radical-producing pathways in the diaphragm include the mitochondria, NADPH oxidase, xanthine oxidase, and/or an increase in reactive iron in the cell. For example, the mitochondria can produce superoxide at complexes I and III along the electron transport chain (7). Also, NADPH oxidase can catalyze the one electron reduction of oxygen into superoxide using NADPH or NADH as the electron donor (8). Moreover, increases in calcium-activated neutral proteases (calpain) can upregulate xanthine oxidase activity, resulting in the formation of excess superoxide production (7). Finally, skeletal muscle atrophy during prolonged periods of immobilization is associated with an increase in free, "reactive" iron (12). Free iron can react with hydrogen peroxide and superoxide to produce the highly reactive hydroxyl radical (7). Determining which of the potential sources of oxidant production is involved in MV-induced oxidative stress is important and could provide a therapeutic intervention to retard MV-induced diaphragmatic contractile dysfunction.
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GRANTS |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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  S. K. Powers, A. N. Kavazis, and J. M. McClung Oxidative stress and disuse muscle atrophy J Appl Physiol, June 1, 2007; 102(6): 2389 - 2397. [Abstract] [Full Text] [PDF]
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