Figures
Abstract
Prolonged muscle exposure to low PO2 conditions may cause oxidative stress resulting in severe muscular injuries. We hypothesize that PO2 cycling preconditioning, which involves brief cycles of diaphragmatic muscle exposure to a low oxygen level (40 Torr) followed by a high oxygen level (550 Torr), can reduce intracellular reactive oxygen species (ROS) as well as attenuate muscle fatigue in mouse diaphragm under low PO2. Accordingly, dihydrofluorescein (a fluorescent probe) was used to monitor muscular ROS production in real time with confocal microscopy during a lower PO2 condition. In the control group with no PO2 cycling, intracellular ROS formation did not appear during the first 15 min of the low PO2 period. However, after 20 min of low PO2, ROS levels increased significantly by ∼30% compared to baseline, and this increase continued until the end of the 30 min low PO2 condition. Conversely, muscles treated with PO2 cycling showed a complete absence of enhanced fluorescence emission throughout the entire low PO2 period. Furthermore, PO2 cycling-treated diaphragm exhibited increased fatigue resistance during prolonged low PO2 period compared to control. Thus, our data suggest that PO2 cycling mitigates diaphragm fatigue during prolonged low PO2. Although the exact mechanism for this protection remains to be elucidated, it is likely that through limiting excessive ROS levels, PO2 cycling initiates ROS-related antioxidant defenses.
Citation: Zuo L, Diaz PT, Chien MT, Roberts WJ, Kishek J, Best TM, et al. (2014) PO2 Cycling Reduces Diaphragm Fatigue by Attenuating ROS Formation. PLoS ONE 9(10): e109884. https://doi.org/10.1371/journal.pone.0109884
Editor: Xiao Su, Chinese Academy of Sciences, China
Received: June 9, 2014; Accepted: September 12, 2014; Published: October 9, 2014
Copyright: © 2014 Zuo et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All relevant data are within the paper and its Supporting Information files.
Funding: Funding provided by Oakland University General Fund G110, Research Excellence Fund of Biomedical Research, and The Ohio State University-Health and Rehabilitation Sciences fund 013000. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Low oxygen/hypoxic conditions can significantly reduce skeletal muscle contraction [1]. In normal resting muscle, it has been reported that skeletal muscles, such as the diaphragm, produce reactive oxygen species (ROS) including hydrogen peroxide (H2O2) [2]. However, when the diaphragm is repetitively stimulated, these muscle fibers generate excessive ROS leading to oxidative stress with accelerated fatigue development [2]. Moreover, the production of ATP is driven by electron transmission through mitochondrial complex I to complex IV, creating a proton gradient across the inner mitochondrial membrane (IMM) and triggering ATP synthesis [3], [4]. Through this mechanism, a small portion of electrons may leak out of the IMM and react with adjacent oxygen molecules to produce superoxide anions, H2O2, and other ROS. Under prolonged low PO2 conditions, the physiological concentration of O2 is altered which results in increased uncoupling between O2 and electron flow, ultimately causing ROS overproduction [5].
A variety of cellular preconditioning pathways associated with muscular protection have been proposed. For instance, ischemic preconditioning (IPC), which consists of ischemic-reperfusion cycles produced by variations in low-high PO2 levels, has been used to prevent cardiac muscle injuries [6]. In addition, IPC initiates intracellular protein kinase pathways, resulting in increased activation of antioxidant enzymes such as catalase [6]. IPC also plays a critical role in protecting the heart against ischemia-reperfusion injuries by opening mitochondrial ATP sensitive potassium channels (mKATP) [7]. The mKATP channels are regulated by several factors, including adenosine, H+, and/or protein kinase C. Thus, these mediators may partially contribute to the protective response involved in preconditioning therapies [8], [9]. Similar to IPC, PO2 cycling preconditioning, which consists of brief periods of lower-higher PO2, significantly protects heart muscle cells subjected to prolonged ischemia by decreasing ROS-induced cell death [7], [10]. In addition, human studies have shown that intermittent low oxygen exposure at low altitude significantly increases an aircraft crew's adaptation to low oxygen conditions experienced at high altitude [11]. Since the method of both IPC and PO2 cycling preconditioning involves brief periods of low and high oxygen levels, it is possible that PO2 cycling follows a similar molecular pathway as IPC. Furthermore, it has been shown that a protocol consisting of PO2 cycling provides a protective response against mesenchymal stem cell (MSC) apoptosis through phosphorylation of extracellular regulated kinase (ERK1/2) and protein kinase B (AKT) [12]. Therefore, it is possible that these signaling factors also may be involved in the molecular mechanism of PO2 cycling in skeletal muscle.
Moreover, lower PO2 or hypoxic conditions may cause changes in the cytosolic redox equilibrium, resulting in a rise in NADPH. The increase subsequently stimulates inositol triphosphate (IP3) receptor mediated release of Ca2+ from the endoplasmic reticulum. This release of Ca2+ activates important cell survival signaling pathways, which may potentially contribute to the preconditioning response during lower PO2 stress [13]. However, the redox mechanism of PO2 cycling preconditioning particularly in respiratory skeletal muscle has not been fully elucidated. The ultimate importance of the work is to develop treatments for those who may experience respiratory muscle fatigue. It is likely that PO2 cycling initiates ROS-related protective responses, particularly in a key muscle of respiration such as the diaphragm, which must be active throughout life [14].
In this study, we tested the hypothesis that PO2 cycling preconditioning decreases intramuscular ROS levels and enhances diaphragm muscle function. Our results demonstrate that PO2 cycling effectively reduces diaphragm fatigue during a prolonged low PO2 (40 Torr) condition, which is accompanied by decreased intracellular ROS levels. These findings provide insight into the molecular redox mechanism of PO2 cycling in diaphragmatic skeletal muscle exposed to a lower PO2 environment.
Materials and Methods
Animals
Male adult C57BL/6 mice (∼20–30 g, average age of ∼3 mo.) were used in accordance with the Ohio State University's and Oakland University's Institutional Laboratory Animal Care and Use Committee (IACUC). We strictly followed the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and Ethics of Animal Experiments. Mice were anesthetized via intraperitoneal (IP) injection with a combination of ketamine (70 mg/kg) and xylazine (10 mg/kg). The diaphragm was quickly removed from the mouse and muscle strips (∼0.5 cm wide, ∼1 cm long, 1–2 strips obtained from each mouse) were dissected from the diaphragm with the corresponding rib attachment and central tendon. After isolation, the muscle strip was kept in Ringer's solution (in mM: 21 NaHCO3, 1.0 MgCl2, 1.2 Na2HPO4, 0.9 Na2SO4, 2.0 CaCl2, 5.9 KCl, 121 NaCl, and 11.5 glucose), at 37°C.
PO2 cycling and muscle function measurement
Function experiments were performed in a contraction chamber (model 800 MS; Danish Myo Technology, Denmark), with the central tendon of the muscle strip sutured to a mobile lever, which was used to adjust the muscle length for optimal performance. The opposite end of the strip was secured to a force transducer (detection range 0–1,600 mN). After being mounted, muscle optimal length (L0) was set as the baseline tension and no adjustments were made throughout the muscle function experiments. All muscles were electrically stimulated (S48 stimulator; Grass Technologies, RI) using square-wave pulses (250-ms train duration, 0.5-ms pulse duration, 70 Hz, 30 V), following previous skeletal muscle function protocols [15], [16]. The A-D board (model ML826; AD instruments, CO) converted the analog signals to digital data, and LabChart 7.3.1 software was used to analyze the function data. The muscle was equilibrated in Ringer's solution for 20–30 min. During the function experiments, the treated muscle strips were switched to a Ringer's solution equilibrated with PO2 of 40 Torr O2 (lower) for 2 min, followed by PO2 of 550 Torr O2 (higher) for 2 min. This PO2 cycle was repeated five times, followed by a prolonged 30-min 40 Torr PO2 period. During this exposure, muscle contractility was evaluated, in order to determine the effect of PO2 cycling on the muscle function. The chamber solution during lower PO2 was found to be 40 Torr and during higher PO2, 550 Torr. In the middle of the 40 Torr PO2 period (from 15–20 min), the muscle was stimulated for 5 min at 37°C and muscle tension development was recorded. The control group followed an identical protocol as the experimental group except for the one intervention of PO2 cycling. Following the removal of the attached rib bone and excess tendon, the diaphragm was first air dried which was followed by a 30-min oven drying. The dry mass was then determined using an analytical balance. To reduce random effects due to animal variance, all function data were normalized by dry weight of the muscle strip (mN/mg).
Regarding the H2O2 treatment with PO2 cycling group for low PO2 contraction measurements, the muscle strips were treated the same as above except that after PO2 cycling, we added H2O2 into the muscle contraction solution. Although H2O2 may degrade rapidly, at sufficient levels, it can enter the cell freely and affect the intracellular activity [17]. Specifically, the muscle was loaded with Ringer's solution with adequate H2O2 (50 µM) for 15 min prior to the 5-min contractions in low PO2 conditions. In addition, the time to reach 50% (T50) of the initial tension in contracting diaphragm muscle during a 5-min low PO2 contraction period was recorded in control, PO2 cycling, and PO2 cycling + H2O2 groups.
The effect of varying H2O2 dosage on muscle tension development was evaluated. The muscle strips were prepared in the same manner as mentioned above. During a high PO2 (550 Torr) period, each muscle strip was equilibrated with Ringer's solution for 15 min followed by a 15 min incubation with a particular dosage of H2O2 (0 µM, 50 µM, 100 µM, 1 mM, to 10 mM, respectively). After incubation, each diaphragm strip was stimulated for 5 min at 37°C and the muscle tension development was recorded.
For the H2O2 treatment group with no PO2 cycling for high PO2 contraction measurements, the muscle strips were exposed to high PO2 (550 Torr). Each muscle strip had two independent 5-min contraction periods, which were separated by a 60-min rest period. The muscle was loaded with H2O2 (50 µM) for 15 min followed by the first 5-min contractile period. The H2O2 was then washed out with fresh Ringer's solution, and the muscle was kept for a 60-min rest period before a second 5-min contraction bout in the absence of H2O2. This protocol was also performed in a blocked order to ensure the statistical value.
Confocal studies
To analyze the effects of PO2 cycling treatment on ROS levels in superfused diaphragm, confocal microscopy was used to measure real-time ROS (H2O2) production in both PO2 cycling-treated and control diaphragm tissue. Specifically, each muscle strip was loaded with a 40 µM solution of dihydrofluorescein diacetate (Hfluor-DA; stock in dimethyl sulfoxide; Sigma-Aldrich) for 30 min. The dye diffused into the intramuscular compartment and was able to chemically react with intracellular ROS (mainly H2O2) resulting in enhanced florescence. For statistical purposes, one muscle strip was taken from each mouse. We used five mice for control and five mice for PO2 cycling treatment. A laser scan confocal microscopy system (Nikon confocal microscope D-Eclipse C1 system) recorded fluorescent emission signals from the tissue sample in a glass bottom culture dish (MatTek Corporation, Ashland, MA) in real time. The treated muscle strips were mounted and superfused with Ringer's solution, followed by PO2 cycling treatment, which was a similar method as described above for the muscle function experiments. To ensure an accurate oxygen level, the chamber was sealed except for the tubing inlets containing gas bubbling and superfusate as well as the temperature probe. The superfusate solution was fully saturated with designated gas and preheated to ensure the temperature in the chamber remained at 37°C. The strips were then subjected to a 10-min baseline period (PO2 of 550 Torr) and a subsequent 30-min 40 Torr PO2 period at 37°C. The control muscles followed the same protocol except that there was no PO2 cycling treatment. During the 40-min experimental period (10 min for baseline of 550 Torr PO2 and 30 min 40 Torr PO2 period), we captured an image (512×512 pixels) every 5 min and calculated the mean fluorescence to determine intramuscular ROS levels. To reduce the signal-to-noise ratio, each recorded image was an average from eight sequentially scanned images within 5 s at each time point. The setup parameters for the confocal imaging system were listed as follows: laser, argon; pinhole: medium or large; excitation, 488 nm; emission, 535±25 nm. The baseline autofluorescence was kept at a minimum and did not interfere with the ROS signal in our set-up. To reduce photobleaching or photodamaging, the laser power was set at ∼15% without noticeably sacrificing image quality. To reduce imaging saturation due to possible excessive ROS in 40 Torr PO2 conditions, the PMT gain was set as low as possible from the start of each experiment. In order to verify that the increased fluorescence signal was due to ROS, a series of antioxidant experiments were conducted. The glutathione peroxidase mimic, ebselen (30 µM), which is an effective ROS scavenger particularly for skeletal muscle tissue [5], was utilized. The animals were divided into 4 experimental groups, including control, PO2 cycling, ebselen, and PO2 cycling + H2O2. In each experimental group, five muscle samples were directly isolated from five fresh isolated diaphragms. For each experiment, we were able to measure ROS levels from ∼8–10 muscle fibers in each image field.
In preliminary studies, we found that the PO2 cycling protocol did not change the fluorescence baseline in the current set-up (data not shown). Each acquired image was analyzed with Adobe Photoshop element 6.0 and the final images were presented in a 300 DPI resolution with LZW compression.
Statistics
By performing the power analysis on the sample, we defined the PO2 cycling effect on the skeletal muscle force development as well as intracellular ROS formation. For instance, we determined the PO2 cycling effect on multiple groups including control, antioxidant (ebselen) treatment and H2O2 application, using the prospective means across these groups. We derived the power when the sample size was ∼5 or greater mice per group by calculating the standard deviation. In addition, data were analyzed using a multi-way ANOVA with the animal as a variable, and expressed as means ± SE. The differences between the two treatments were statistically determined by a series of post-ANOVA contrast analyses using JMP software (SAS Institute, Cary, NC). Specifically, the post-ANOVA contrasts involve the comparison among all the groups of subjects and the display of the statistical difference between each pair of data. The treated groups, such as PO2 cycling, antioxidant (ebselen) treatment and H2O2 application, were used to compare with the control group, revealing any potential significance. P<0.05 was regarded to be significant.
Results
Representative confocal images of the same muscular area in each group are illustrated in Fig. 1. Fluorescence (green color), which represents ROS levels, increased substantially at the end of prolonged 40 Torr PO2 period (30 min, Fig. 1B) compared to baseline in the control group (Fig. 1A). However, after PO2 cycling treatment, fluorescence displayed no significant change at the end of 40 Torr PO2 (Fig. 1D) when compared to baseline (Fig. 1C). The disappearance of increased fluorescence emission in PO2 cycling treatment demonstrated that PO2 cycling effectively suppressed 40 Torr PO2-induced intracellular ROS levels in the diaphragm. In addition, ebselen treated muscle strips showed no fluorescence increase at the end of 40 Torr PO2 exposure (Fig. 1F). Interestingly, exogenous addition of H2O2 (50 µM) mitigated the antioxidant effect of PO2 cycling (Fig. 1H).
A: control muscle before 40 Torr PO2. B: the same area of A at the end of 40 Torr PO2. C: PO2 cycling treated muscle before 40 Torr PO2. D: the same area of C at the end of 40 Torr PO2. E: Ebselen (Eb) treated muscle before 40 Torr PO2. F: the same area of E at the end of 40 Torr PO2. G: PO2 cycling + H2O2 treated muscle before 40 Torr PO2. H: the same area of G at the end of 40 Torr PO2.
Grouped data of mean florescence during 40 Torr PO2 are illustrated in Fig. 2A. At the end of 40 Torr PO2 periods, ROS levels were elevated from baseline in the control group. However, in the PO2 cycling group, ROS levels were kept low compared to control (1.00±0.04 RU vs. 1.48±0.05 RU; n = 5 from five animals; P<0.05). Furthermore, in the control group, intracellular ROS elevation did not appear within the first 15 min of 40 Torr PO2 period. After 20 min, ROS levels were enhanced and these increases lasted until the end of the 30 min period. Fig. 2B displayed the intracellular ROS fluorescence rate (RU/min). In the control group, this rate was close to zero for the first 15 min of 40 Torr PO2, followed by three ROS bursts occurring at 20, 25, and 30 min, represented by positive values shown in Fig. 2B (in RU/min). The first ROS burst was relatively smaller (∼50% less) compared to the other two larger bursts. However, in PO2 cycling-treated diaphragm muscles, no ROS burst occurred at 40 Torr PO2. It is important to note that although Fig. 2B looks similar to Fig. 2A, they refer to two separate measurements: fluorescence in Fig. 2A and fluorescence rate in Fig. 2B. In other words, Fig. 2B shows the mathematical slope of the fluorescence increase/decrease, indicating how fast the signal changes while Fig. 2A shows the level or the intensity of fluorescence.
A: averaged fluorescence data recorded in a relative unit (RU). Data showed intracellular ROS levels from control, PO2 cycling, ebselen (Eb), and PO2 cycling + H2O2 treated diaphragm muscle (*significantly different from PO2 cycling, P<0.05; #significantly different from Eb treatment, P<0.05; †significantly different from control, P<0.05). B: intracellular ROS burst represented by fluorescence rate. Data was recorded in a relative unit per min (RU/min) from control, PO2 cycling, Eb, and PO2 cycling + H2O2 (50 µM) treated diaphragm muscle under 40 Torr PO2. Fluorescence data was recorded every 5 min (*significantly different from PO2 cycling, P<0.05; #significantly different from Eb treatment, P<0.05; †significantly different from control, P<0.05).
Muscle absolute tension data during 40 Torr PO2 are shown in Fig. 3. In the control group, the tension (in mN/mg muscle dry weight) at 0 min and the tension at each subsequent time point thereafter (1–5 min), was significantly lower than the PO2 cycling group (n = 5 from five animals, P<0.05), suggesting that PO2 cycling ameliorated skeletal muscle resistance to fatigue during the 40 Torr PO2 period (Fig. 3A). The tension decline rate in PO2 cycling treated muscles markedly slowed down in the first 3 min compared to control (n = 5 from five animals for each group, P<0.05, Fig. 3B). However, after 3 min the decline rate of all groups was similar, indicating that PO2 cycling had no effect on the force decline rate for later fatigue development. Furthermore, the time to 50% of the initial tension (T50) from PO2 cycling diaphragm muscle was significantly prolonged when compared to control diaphragm (in seconds, 216.2±33.0 vs. 99.5±10.0; n = 5 from five animals, P<0.05, Fig. 4). However, this difference disappeared in the presence of H2O2 in the PO2 cycling + H2O2 group (in seconds, 117.1±7.2 vs. 99.5±10.0; n = 5 from five animals, Fig. 4). Maximal diaphragm force was always measured prior to low PO2 exposure. The corresponding muscle absolute tension values were reported in Table 1.
A: absolute tension (mN/mg) was recorded for 5 min from control, PO2 cycling, and PO2 cycling + H2O2 group (*significantly different from PO2 cycling, P<0.05). B: data showing the tension decline rate (RU/min) from control, PO2 cycling, and PO2 cycling + H2O2 (50 µM) muscles during a 5-min contractile period under 40 Torr PO2 (*significantly different from PO2 cycling, P<0.05; #significantly different from PO2 cycling + H2O2, P<0.05).
A: a typical chart record illustrating the duration of T50 in a contracting diaphragm muscle during a 5-min contraction period. B: summarized T50 values from control, PO2 cycling, and PO2 cycling + H2O2 (50 µM) groups during the contraction (*significantly different from control, P<0.05; #significantly different from PO2 cycling + H2O2, P<0.05).
We observed that ebselen completely quenched the 40 Torr PO2-induced ROS signal in the control (Fig. 1E and F, Fig. 2; n = 4), which demonstrated a similar effect to the PO2 cycling treatment group (Fig. 1C and D, Fig. 2, n = 5). The addition of a small amount of H2O2 (50 µM) entirely abolished the PO2 cycling-induced ROS inhibition effect in the confocal experiments (Fig. 1G and H). Grouped data are shown in Fig. 2A. Following 15 min from the onset of 40 Torr PO2 period, fluorescence was significantly higher in PO2 cycling + H2O2 group than the control, PO2 cycling, and ebselen groups. However, after 30 min of 40 Torr PO2 period, the control group showed higher fluorescence than the PO2 cycling + H2O2 and ebselen groups, respectively (n = 5, P<0.05). In Fig. 2B, at 15 min during 40 Torr PO2 periods, there was a larger fluorescent burst in the PO2 cycling + H2O2 treatment group compared to the other groups. A large fluorescent burst in the control group occurred ∼10 min later than in the PO2 cycling + H2O2, while there were no bursts in ebselen treatment groups, respectively (n = 5, P<0.05).
Furthermore, under high PO2 conditions (550 Torr), we investigated the effect of a small amount of H2O2 (50 µM) on muscle contraction as shown in Fig. 5 (representative curves) and Fig. 6 (grouped data). The tension development (mN/mg) and the tension decline rate (RU/min) at 1–5 min during the 5-min contraction were recorded in the presence or absence of H2O2. Both figures clearly illustrate that H2O2 had no marked effect on muscle function at a level of 50 µM (n = 6).
A: H2O2 was added 15 min prior to the first 5-min contractile period followed by a H2O2 washout and 60 min rest period before the second 5- min contractile period in the absence of H2O2. B: The first 5-min contraction in the absence of H2O2 was followed by the second 5-min contractile period in the presence of H2O2.
A: Tension development (mN/mg) at 1–5 min during the 5-min contraction period in the presence vs. absence of H2O2. B: Tension decline rate (RU/min) during the 5- min contraction period at 1–5 min in the presence vs. absence of H2O2.
Moreover, H2O2 dosage experiments were performed in a range from 0 µM to 10 mM under high PO2 conditions (550 Torr) as shown in Fig. 7. Muscle tension development was recorded for the maximal contraction during the baseline period and the initial and end contractions during the 5-min contraction period. There was no significant difference between the control group (0 µM) and the 50 µM group. However, the 100 µM and 1 mM groups (n = 9 for both groups) did show a significant decrease in muscle function during the end contraction when compared to both the control and 50 µM groups (P<0.05). The greatest declines in muscle tension were observed in the 10 mM group (n = 8) as both the initial and end contractions showed a marked decrease in muscle tension development in comparison to all other dosage groups (P<0.05).
Data showing the muscle tension development (mN/mg) during the H2O2 dosage treatments in mouse diaphragm strips [0 µM (control), 50 µM, 100 µM, 1 mM and 10 mM] in high PO2 (550 Torr). *Significantly different from control (P<0.05). #Significantly different from 50 µM H2O2 (P<0.05). †Significantly different from 100 µM H2O2 (P<0.05). ‡Significantly different from 1 mM H2O2 (P<0.05).
Discussion
The current study provides evidence that the PO2 cycling preconditioning procedure we used reduces intracellular ROS levels in respiratory skeletal muscle during prolonged low PO2. The absolute skeletal muscle tension and T50 were both greater in the PO2 cycling group than the control group, but the addition of a small amount of ROS (H2O2) reduced these values to control levels. However, this amount of ROS was so marginal that it exerted no significant effect on muscle function during hyperoxia. Collectively, these data indicate that the protection of PO2 cycling on the diaphragm is related to the reduced levels of intracellular ROS signaling molecules.
Dihydrofluorescein (Hfluor) is a highly sensitive intracellular probe commonly used for fluorescent detection of ROS. Fluorescein (Fluor) formation results when Hfluor reacts with ROS [5]. Previous research has shown that Hfluor is much less sensitive to nitric oxide (NO) compared to its analog dichlorfluorescein (DCFH) and also shows a higher resistance to photobleaching than DCFH [5], [18], [19]. Since it is superior for detecting ROS (particularly H2O2), it was used in our experiments. Our results showed that there was no marked ROS formation in the muscle during the first 15 min of a 40 Torr PO2 period. In the control group (Fig. 2A), ROS levels were significantly increased after 20 min from the initiation of 40 Torr PO2, which may suggest that during this timeframe antioxidant defenses were overwhelmed in diaphragm muscle not treated with PO2 cycling. Moreover, the completely abolished ROS signals in the antioxidant (ebselen) treated control group (Fig. 1 and 2), confirms the existence of ROS, which seems to be quenched by PO2 cycling treatment in our experiments (Fig. 1 and 2). Interestingly, after PO2 cycling treatment, extracellular addition of a small amount of ROS (50 µM H2O2), which has no effect on normal muscle function (Fig. 5 and 6), completely negated the PO2 cycling effect. These observations suggest to us that PO2 cycling may be involved in the initiation of intracellular antioxidant signaling pathways.
It should be noted that the experimental conditions used to detect ROS are different from those used to evaluate skeletal muscle function in our settings. ROS detection was performed in unstimulated muscle, while function was assessed in contracting muscle, for the following reasons: 1) Due to large motion artifact, it is extremely difficult to perform muscle function experiments under confocal microscopy; 2) The muscle function experiment is focused on measurement of maximal force and time to fatigue. However, the confocal experiment is primarily designed to determine intracellular ROS levels in the muscle.
Evidence has shown that PO2 cycling triggers the expression of superoxide dismutase (SOD), an endogenous antioxidant, which may further contribute to reduced ROS levels [7], [10], [20]. This is highly consistent with our observations of reduced ROS level in PO2 cycling treated skeletal muscle. Similar to heart studies which show that both IPC and PO2 cycling are mediated by ROS [7], intracellular ROS levels were also critical for PO2 cycling efficacy in diaphragm muscle. There is a potential concern that PO2 cycling could have altered mitochondrial function and integrity. However, our muscle function data (Fig. 3 and 4) suggest that fatigue resistance in PO2 cycling treated mouse diaphragm muscle was substantially greater than that of control muscle. Thus, it is likely that mitochondrial activity was not negatively altered by PO2 cycling treatment. Although in the current study design we are unable to determine whether PO2 cycling treatment causes decreased ROS production or increased antioxidant scavenging, it is likely that there is a specific redox mechanism that suppresses ROS generation during prolonged respiratory muscle exposure to the 40 Torr PO2. The detailed mechanism associated with PO2 cycling protection, however, is still unclear and requires further study. In addition, based on a similar previous study of PO2 levels in myocytes, 40–550 Torr PO2 was an effective setting to initiate intramuscular redox changes in skeletal muscle [21], [22].
Interestingly, we noticed that at 15 min during the 40 Torr PO2 period, ROS levels were significantly higher in the PO2 cycling + H2O2 group compared to the other groups; yet, at 30 min, the control group showed a higher ROS level than the PO2 cycling + H2O2 group (Fig. 2A). This suggests that the two treatment plans stimulate a time-dependent intracellular ROS formation mechanism. In addition, we evaluated the ROS generation rate, defined as a ROS burst and represented by the fluorescence rate, as shown in Fig. 2B. H2O2 addition after PO2 cycling treatment induced the first ROS burst at 15 min under the 40 Torr PO2 conditions, which occurred ∼10 min earlier than a large ROS burst in the control group. Collectively, these observations suggest a faster diffusion of extracellular ROS (H2O2) into the intramuscular compartment, compared to the intracellular ROS generation in the control muscle. However at a later time (after 25 min during low PO2), the control muscle showed a higher ROS formation rate than the PO2 cycling + H2O2 group. This could be due to leakage of H2O2 into the perfusate. Nevertheless, these data further demonstrate a potential antioxidant-like effect exerted by PO2 cycling, and this effect can be disturbed by a small addition of ROS, which was not sufficient to influence muscle contractility (Fig. 5 and 6).
Moreover, it is suggested that PO2 cycling mitigates fatigue within the diagram during hypoxia. Although the exact mechanism by which this occurs is unclear, it is likely that ROS play a significant role. Accordingly, our data (Fig. 1 and 2) suggest that low levels of ROS negate the benefits of PO2 cycling and may be involved in other signaling events, including antioxidant cascades. Figures 5 and 6 illustrate the relationship between ROS level and muscular force generation. These results are supported by previous studies in which it was concluded that low levels of H2O2 may work more towards signaling pathways since they do not directly impact force generation in the muscle [23], [24]. Further research into the effect of both ROS and PO2 cycling on diaphragm force generation and related muscular mechanisms may lead to potential therapies to mitigate muscle fatigue during hypoxia.
The H2O2 dosage experiments (Fig. 7) on the diaphragm function suggest that high levels of ROS (H2O2) such as at 1 mM or 10 mM levels markedly reduce muscle function; however, lower levels of ROS, such as 50 µM, do not compromise muscle function as compared to control. In addition, our confocal experiments have clearly shown that this low level of ROS (50 µM) minimized the PO2 cycling effect confirming that rather than damaging muscle directly, low levels of ROS may be a potential mediator for the signaling events involved in PO2 cycling preconditioning.
There are a number of intracellular sources of ROS in skeletal muscle, including the mitochondria, xanthine oxidase (XO), peroxisomes, and NADPH oxidase [25]. For example, under respiratory stress, such as ischemia or low PO2/hypoxia, xanthine dehydrogenase converts to XO, which is subsequently released into circulation and produces ROS [22], [25]. In skeletal muscle myocytes, NADPH oxidase is another likely candidate for ROS formation during hypoxic stress during injuries [23]. Additionally, the mitochondria produce low levels of superoxide anion under normal conditions [26], but in the lack of oxygen, the mitochondria experience excessive oxidant production [5], [26], [27]. Accordingly in such a condition, the mitochondria function as a source of increased level of superoxide, one of the common precursors to most ROS. This increase in intracellular ROS could potentially overwhelm natural antioxidant defense systems, leading to impaired muscle function [2], [28]–[30]. Our data (Fig. 2B) showed that three ROS oxidative bursts (represented by positive rate values) occurred after 15 min of the 40 Torr period in the control muscle, which may indicate that intracellular antioxidant defenses have a 15 min effective period until they are eventually negated by subsequent ROS formation. This timeframe is within the regular activation time range of mitochondrial antioxidant enzymes [30], [31]. Therefore, both XO and mitochondria are possible sources of ROS in this study.
Moreover, our findings have shown considerable evidence regarding the protective effects of PO2 cycling training on skeletal muscle function (Fig. 3A). During the middle of the 40 Torr PO2 period, the initial tension of the PO2 cycling treated muscle strips was higher than that of control, and this trend kept increasing to ∼4 fold greater than control from 1- to 5-min suggesting that PO2 cycling progressively alleviated muscle fatigue. However, after H2O2 was loaded into the muscle at a relatively low dosage (Fig. 5 and 6), the protection from PO2 cycling was diminished. This supports that the ROS signaling molecules may possibly play a negative role in the PO2 cycling mechanism. PO2 cycling markedly reduced the tension decline rate at both 1-, 2- and 3-min from the initial contraction compared to control. These rate differences were minimized at 4- and 5-min, respectively. The addition of H2O2 also significantly interrupted the PO2 cycling effect on the tension decline rate at 2- and 3-min in the middle of the 40 Torr PO2 period (Fig. 3B), confirming that ROS signaling molecules adversely affect PO2 cycling pathways. A possible correlation among PO2 cycling, muscle fatigue and ROS implies that PO2 cycling is able to boost muscle contractility during fatigue, which is consistent with Clanton's and Reid's results [1], [32]. Specifically, PO2 cycling could result in a gradual increase in the production of endogenous antioxidant enzymes, allocating the additional antioxidants to serve as a “reservoir” that can be promptly accessed in response to spontaneous exposure to stressful conditions. This idea is consistent with previous research suggesting that PO2 cycling significantly increases the expression level of intramuscular antioxidants such as SOD [7].
Previous research has shown that the intramuscular PO2 during strenuous exercise is ∼4 Torr, while ∼30 Torr conditions are seen in resting muscle [33]. In the current research, we created a relatively lower PO2 condition by equilibrating 40 Torr PO2 and a hyperoxic condition with 550 Torr based on previous studies in skeletal muscle [5], [15], [21], [34]. However, the intracellular PO2 was difficult to determine under our experimental set-up, particularly in a contracting muscle with marked motion. During exhaustive exercise, skeletal muscle conditions fluctuate between higher PO2 and lower PO2, which frequently occur especially during high intensity interval trainings. This cycling in intracellular oxygen exerts an intrinsic preconditioning effect, similar to the PO2 cycling protocol implemented in our studies. Additionally, it has been shown that increased levels of catalase and SOD are expressed in skeletal muscle during exercise training, resulting in a reduction of ROS level and oxidative stress [35]–[38]. Similar to exercise training, PO2 cycling therapy may be an alternative method for increasing muscular endurance. Moreover, it is worth noting that a small amount of ROS (50 µM H2O2) has no effect on normal muscle function (Fig. 5 and 6). However, this dosage completely abolished the PO2 cycling protection in low PO2 conditions (Fig. 3). The exact mechanism of this response is still unknown, which will be an area of future studies.
Some limitations appear in our study. First, we are unable to determine whether 40 Torr in the solution can cause intramuscular hypoxia. Second, it is possible that our PO2 cycling protocol, by itself, can directly affect intracellular ROS and muscle fatigue without additional mediators. Third, it is difficult to measure the exact intracellular PO2 level in a functioning diaphragm. This is mainly because of a marked O2 diffusion gradient across the multiple layers of diaphragm tissue. Although previous research has shown that in a similar condition to 40 Torr PO2, intracellular levels of NADH in the diaphragm significantly increase [5], it is not clear whether 40 Torr PO2 in the superfusate can cause the intracellular compartments hypoxic. Lastly, since O2 cannot transport across the different layers equally in the whole muscle, it is more likely that a hypoxic condition may occur in the core of the muscle than the peripheral region [39]. In addition, intracellular PO2 in skeletal muscle is ∼10 Torr at rest, but it quickly drops to 3–5 Torr during intense exercise [40]. It is likely that the transition between 550 Torr to 40 Torr triggers mismatches of oxygen supply to the diaphragm, which may be sufficient to induce a transient ROS formation as described in our earlier research [5]. Our study demonstrated that intracellular ROS is elevated in single myofibers during a similar PO2 condition [21]. Interestingly, this level of oxygen (3–5 Torr) is regarded as normal for exercising human muscles [33]. Precisely controlling the intramuscular O2 condition within the whole muscle preparation is highly challenging and therefore should be the focus of future research.
Perspectives and Significance
This study demonstrates that PO2 cycling mediates beneficial responses through reducing intracellular ROS levels in respiratory muscle. PO2 cycling is a drug-free treatment that possibly stimulates the diaphragm to activate its own antioxidant defense systems to resist fatigue development. This may be an effective method for enhancing muscular endurance. In addition, the current in vitro study provides a redox perspective into mouse respiratory muscle under optimal preconditions.
Supporting Information
Dataset S1.
ROS fluorescence (figures 1 and 2).
https://doi.org/10.1371/journal.pone.0109884.s001
(XLSX)
Dataset S2.
Muscle tension and tension decline rate (figure 3).
https://doi.org/10.1371/journal.pone.0109884.s002
(XLSX)
Dataset S3.
Time to reach 50% (T50) of the initial tension (figure 4).
https://doi.org/10.1371/journal.pone.0109884.s003
(XLSX)
Dataset S4.
Representative H2O2 (50 µM) contraction curves (figure 5).
https://doi.org/10.1371/journal.pone.0109884.s004
(XLSX)
Dataset S5.
Grouped H2O2 (50 µM) data (figure 6).
https://doi.org/10.1371/journal.pone.0109884.s005
(XLSX)
Dataset S6.
Grouped data of varying H2O2 dosage (figure 7).
https://doi.org/10.1371/journal.pone.0109884.s006
(XLSX)
Acknowledgments
We thank Dr. Lan Jiang, Benjamin Pannell, Alexander C. Ziegler, Anthony Re, Shenali Wickramanayake, Tingyang Zhou and Julia Stimpfl for their assistance.
Author Contributions
Conceived and designed the experiments: LZ PTD MTC WJR JK TMB PDW. Performed the experiments: LZ MTC WJR. Analyzed the data: LZ WJR. Contributed reagents/materials/analysis tools: LZ WJR. Wrote the paper: LZ PTD MTC WJR JK TMB PDW. Designed the method of function measurement: LZ. Design of the manuscript writing style: LZ PDW.
References
- 1. Mohanraj P, Merola AJ, Wright VP, Clanton TL (1998) Antioxidants protect rat diaphragmatic muscle function under hypoxic conditions. J Appl Physiol 84: 1960–1966.
- 2. Reid MB, Haack KE, Franchek KM, Valberg PA, Kobzik L, et al. (1992) Reactive oxygen in skeletal muscle. I. Intracellular oxidant kinetics and fatigue in vitro. J Appl Physiol 73: 1797–1804.
- 3. Semenza GL (2011) Hypoxia-inducible factor 1: regulator of mitochondrial metabolism and mediator of ischemic preconditioning. Biochim Biophys Acta 1813: 1263–1268.
- 4. Chance B, Sies H, Boveris A (1979) Hydroperoxide metabolism in mammalian organs. Physiol Rev 59: 527–605.
- 5. Zuo L, Clanton TL (2005) Reactive oxygen species formation in the transition to hypoxia in skeletal muscle. Am J Physiol Cell Physiol 289: C207–216.
- 6. Choi YS, Cho KO, Kim EJ, Sung KW, Kim SY (2007) Ischemic preconditioning in the rat hippocampus increases antioxidant activities but does not affect the level of hydroxyl radicals during subsequent severe ischemia. Exp Mol Med 39: 556–563.
- 7. Zuo L, Roberts WJ, Tolomello RC, Goins AT (2012) Ischemic and hypoxic preconditioning protect cardiac muscles via intracellular ROS signaling. Front Biol 8: 305–311.
- 8. Gross GJ, Auchampach JA (1992) Blockade of ATP-sensitive potassium channels prevents myocardial preconditioning in dogs. Circ Res 70: 223–233.
- 9. Ravingerova T, Lokebo JE, Munch-Ellingsen J, Sundset R, Tande P, et al. (1998) Mechanism of hypoxic preconditioning in guinea pig papillary muscles. Mol Cell Biochem 186: 53–60.
- 10. Vanden Hoek TL, Becke LB, Shao Z, Li C, Schumacker PT (1998) Reactive oxygen species released from mitochondria during brief hypoxia induced preconditioning in cardiomyocytes. J Biol Chem 273: 18092–18098.
- 11. Muza SR (2007) Military applications of hypoxic training for high-altitude operations. Med Sci Sports Exerc 39: 1625–1631.
- 12. Wang JA, Chen TL, Jiang J, Shi H, Gui C, et al. (2008) Hypoxic preconditioning attenuates hypoxia/reoxygenation-induced apoptosis in mesenchymal stem cells. Acta Pharmacol Sin 29: 74–82.
- 13. Bickler PE, Fahlman CS, Gray J, McKleroy W (2009) Inositol 1,4,5-triphosphate receptors and NAD(P)H mediate Ca2+ signaling required for hypoxic preconditioning of hippocampal neurons. Neuroscience 160: 51–60.
- 14.
Zuo L, Hallman AH, Yousif MK, Chien MT (2012) Oxidative stress, respiratory muscle dysfunction, and potential therapeutics in chronic obstructive pulmonary disease. DOI: 101007/s11515-012-1251-x Front Biol 7: : 506–513.
- 15. Zuo L, Nogueira L, Hogan MC (2011) Effect of pulmonary TNF-alpha overexpression on mouse isolated skeletal muscle function. Am J Physiol Regul Integr Comp Physiol 301: R1025–1031.
- 16. Roberts WJ, Zuo L (2012) Hypoxic preconditioning reduces reoxygenation injuries via PI3K in respiratory muscle. GSTF JBio 2: 42–53.
- 17. Henriksen EJ (2013) Effects of H2O2 on insulin signaling the glucose transport system in mammalian skeletal muscle. Methods Enzymol 528: 269–278.
- 18. Zuo L, Clanton TL (2002) Detection of reactive oxygen and nitrogen species in tissues using redox-sensitive fluorescent probes. Methods Enzymol 352: 307–325.
- 19. Hempel SL, Buettner GR, O'Malley YQ, Wessels DA, Flaherty DM (1999) Dihydrofluorescein diacetate is superior for detecting intracellular oxidants: comparison with 2′,7′-dichlorodihydrofluorescein diacetate, 5(and 6)-carboxy-2′,7′-dichlorodihydrofluorescein diacetate, and dihydrorhodamine 123. Free Radic Biol Med 27: 146–159.
- 20. Chen CF, Tsai SY, Ma MC, Wu MS (2003) Hypoxic preconditioning enhances renal superoxide dismutase levels in rats. J Physiol 552: 561–569.
- 21. Zuo L, Shiah A, Roberts WJ, Chien MT, Wagner PD, et al. (2013) Low Po(2) conditions induce reactive oxygen species formation during contractions in single skeletal muscle fibers. Am J Physiol Regul Integr Comp Physiol 304: R1009–1016.
- 22. Zuo L, Nogueira L, Hogan MC (2011) Reactive oxygen species formation during tetanic contractions in single isolated Xenopus myofibers. J Appl Physiol 111: 898–904.
- 23. Brotto MA, Nosek TM (1996) Hydrogen peroxide disrupts Ca2+ release from the sarcoplasmic reticulum of rat skeletal muscle fibers. J Appl Physiol (1985) 81: 731–737.
- 24. Penheiter AR, Bogoger M, Ellison PA, Oswald B, Perkins WJ, et al. (2007) H(2)O(2)-induced kinetic and chemical modifications of smooth muscle myosin: correlation to effects of H(2)O(2) on airway smooth muscle. J Biol Chem 282: 4336–4344.
- 25. Moylan JS, Reid MB (2007) Oxidative stress, chronic disease, and muscle wasting. Muscle Nerve 35: 411–429.
- 26. Raedschelders K, Ansley DM, Chen DD (2012) The cellular and molecular origin of reactive oxygen species generation during myocardial ischemia and reperfusion. Pharmacol Ther 133: 230–255.
- 27. Zhang L, Yu L, Yu CA (1998) Generation of superoxide anion by succinate-cytochrome c reductase from bovine heart mitochondria. J Biol Chem 273: 33972–33976.
- 28. Andrade FH, Reid MB, Allen DG, Westerblad H (1998) Effect of hydrogen peroxide and dithiothreitol on contractile function of single skeletal muscle fibres from the mouse. J Physiol 509 (Pt 2): 565–575.
- 29. MacFarlane NG, Miller DJ (1992) Depression of peak force without altering calcium sensitivity by the superoxide anion in chemically skinned cardiac muscle of rat. Circ Res 70: 1217–1224.
- 30. Jezek P, Hlavata L (2005) Mitochondria in homeostasis of reactive oxygen species in cell, tissues, and organism. Int J Biochem Cell Biol 37: 2478–2503.
- 31. Zuo L, Christofi FL, Wright VP, Liu CY, Merola AJ, et al. (2000) Intra- and extracellular measurement of reactive oxygen species produced during heat stress in diaphragm muscle. Am J Physiol Cell Physiol 279: C1058–1066.
- 32. Reid MB (2008) Free radicals and muscle fatigue: Of ROS, canaries, and the IOC. Free Radic Biol Med 44: 169–179.
- 33. Wagner PD (2012) Muscle intracellular oxygenation during exercise: optimization for oxygen transport, metabolism, and adaptive change. Eur J Appl Physiol 112: 1–8.
- 34. Zuo L, Hallman AH, Roberts WJ, Wagner PD, Hogan MC (2014) Superoxide release from contracting skeletal muscle in pulmonary TNF-alpha overexpression mice. Am J Physiol Regul Integr Comp Physiol 306: R75–81.
- 35. Caldarera CM, Guarnieri C, Lazzari F (1973) Catalase and peroxidase activity of cardiac muscle. Boll Soc Ital Biol Sper 49: 72–77.
- 36. Jenkins RR, Friedland R, Howald H (1984) The relationship of oxygen uptake to superoxide dismutase and catalase activity in human skeletal muscle. Int J Sports Med 5: 11–14.
- 37. Sen CK (1995) Oxidants and antioxidants in exercise. J Appl Physiol 79: 675–686.
- 38. Criswell D, Powers S, Dodd S, Lawler J, Edwards W, et al. (1993) High intensity training-induced changes in skeletal muscle antioxidant enzyme activity. Med Sci Sports Exerc 25: 1135–1140.
- 39. Barclay CJ (2005) Modelling diffusive O(2) supply to isolated preparations of mammalian skeletal and cardiac muscle. J Muscle Res Cell Motil 26: 225–235.
- 40. Richardson RS, Noyszewski EA, Kendrick KF, Leigh JS, Wagner PD (1995) Myoglobin O2 desaturation during exercise. Evidence of limited O2 transport. J Clin Invest 96: 1916–1926.