Journal of Applied Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

J Appl Physiol 98: 1753-1760, 2005. First published January 7, 2005; doi:10.1152/japplphysiol.01022.2004
8750-7587/05 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
98/5/1753    most recent
01022.2004v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (9)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wright, V. P.
Right arrow Articles by Clanton, T. L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wright, V. P.
Right arrow Articles by Clanton, T. L.

Superoxide scavengers augment contractile but not energetic responses to hypoxia in rat diaphragm

V. P. Wright,1,2 P. F. Klawitter,1,3 D. F. Iscru,1,2 A. J. Merola,1,4 and T. L. Clanton1,2

1The Dorothy M. Davis Heart and Lung Research Institute, and Departments of 2Internal Medicine (Division of Pulmonary, Critical Care and Sleep Medicine), 3Emergency Medicine, and 4Molecular and Cellular Biochemistry, The Ohio State University, Columbus, Ohio

Submitted 16 September 2004 ; accepted in final form 17 December 2004


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Acute exposure to severe hypoxia depresses contractile function and induces adaptations in skeletal muscle that are only partially understood. Previous studies have demonstrated that antioxidants (AOXs) given during hypoxia partially protect contractile function, but this has not been a universal finding. This study confirms that specific AOXs, known to act primarily as superoxide scavengers, protect contractile function in severe hypoxia. Furthermore, the hypothesis is tested that the mechanism of protection involves preservation of high-energy phosphates (ATP, creatine phosphate) and reductions of Pi. Rat diaphragm muscle strips were treated with AOXs and subjected to 30 min of hypoxia. Contractile function was examined by using twitch and tetanic stimulations and the degree of elevation in passive force occurring during hypoxia (contracture). High-energy phosphates were measured at the end of 30-min hypoxia exposure. Treatment with the superoxide scavengers 4,5-dihydroxy-1,3-benzenedisulfonic acid (Tiron, 10 mM) or Mn(III)tetrakis(1-methyl-4-pyridyl) porphyrin pentachloride (50 µM) suppressed contracture during hypoxia and protected maximum tetanic force. N-acetylcysteine (10 or 18 mM) had no influence on tetanic force production. Contracture during hypoxia without AOXs was also shown to be dependent on the extracellular Ca2+ concentration. Although hypoxia resulted in only small reductions in ATP concentration, creatine phosphate concentration was decreased to ~10% of control. There were no consistent influences of the AOX treatments on high-energy phosphates during hypoxia. The results demonstrate that superoxide scavengers can protect contractile function and reduce contracture in hypoxia through a mechanism that does not involve preservation of high-energy phosphates.

skeletal muscle; 4,5-dihydroxy-1,3-benzenedisulfonic acid; N-acetylcysteine; Mn(III)tetrakis(1-methyl-4-pyridyl) porphyrin pentachloride; calcium regulation

ACUTE EXPOSURE TO SEVERE HYPOXIA depresses contractile function and induces adaptations in skeletal muscle by mechanisms that are only partially understood. One well-known mechanism involves the influence of the buildup of energy metabolites such as inorganic phosphate (Pi) and H+, which can provide inhibitory influences on force development and Ca2+ sensitivity (20). A previous study demonstrated that antioxidant treatments during brief periods of severe hypoxia in rat diaphragm partially preserve contractile function, both during and after exposure to hypoxia, suggesting that altered redox balance may also play a role in hypoxia (34).

In this study, the hypothesis was tested that the mechanism by which antioxidants (AOXs) influence contractile function in hypoxia is through preservation of high-energy phosphates [ATP and creatine phosphate (CrP)] and reduction of Pi. There is considerable rationale for this hypothesis, because glucose transport, glycolysis (5), and oxidative phosphorylation (26) have all been shown to be inhibited by exposure to elevated levels of oxidants. Furthermore, oxidants such as peroxynitrite (ONOO–) appear to inhibit glycolysis in heart muscle during ischemia (10, 11). In hearts treated with the superoxide (O2–·) scavenger Tiron, CrP levels are preserved in rat cardiac muscle at the end of an ischemic exposure (25). The hypothesis was addressed in this study by measuring high-energy phosphates, at the end of a 30-min exposure to severe hypoxia in the presence or absence of various AOXs.

Another purpose of this study was to evaluate the influence of antioxidant treatment on the phenomenon of "contracture," a term used to describe the gradual increase in passive force during exposure to severe hypoxia. Contracture in skeletal muscle is modest compared with cardiac muscle but resembles behavior similar to that seen in ischemic heart, where it is believed to reflect responses to elevated free sarcomeric Ca2+ (22).


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
General procedures.   All animal procedures were performed in accordance with The Ohio State University Institutional Lab Animal Care and Use Committee Guidelines. Male Sprague-Dawley rats (360–481 g) were anesthetized with ketamine (100 mg/kg) and xylazine (20 mg/kg), tracheotomized, and mechanically ventilated. The whole diaphragm was excised and immediately placed in oxygenated (95% O2-5% CO2) Ringer salt solution (in meq/l: 21 NaHCO3, 0.9 NaSO4, 1.2 Na2HPO4, 1.0 MgCl2, 2.0 CaCl2, 5.9 KCl, and 121 NaCl with 2.07 g/l glucose and 10 µM D-tubocurarine). Four diaphragm strips from the costal diaphragm were cut, ~5 mm in width, with portions of the central tendon and ribs attached. The strips were mounted vertically in four tissue baths with oxygenated Ringer's solution and maintained at 37°C. Optimal length and maximum field stimulation current were determined for each strip by standard methods (34). Strips were thermo-equilibrated for 15 min while being intermittently stimulated with one twitch every 20 s. Then initial, untreated force-frequency relationships were determined by using a twitch stimulation and stimulations at 20, 30, 40, 50, 60, 80, 100, and 150 Hz, each maintained for 400 ms, with a 20-s recovery interval between stimulations. Before hypoxia, preloads (passive tensions) were reset, when necessary, to the predetermined level obtained at baseline optimal length determination (Fig. 1). Once the tissues were exposed to hypoxia, there was no adjustment of passive force, because it was one of the measured variables.



View larger version (9K):
[in this window]
[in a new window]
  		Fig. 1. Schematic of functional protocol. Arrows indicate when preloads were adjusted. AOX, antioxidant; FF, force-frequency measurement; Lo, optimum length determination, Imax, maximum field current.

 
Drug effects.   After initial contractile measurements, bathing solutions were changed to fresh, fully oxygenated, 37°C Ringer's buffer, with or without AOXs. The AOXs utilized included 1) 10 mM Tiron (4,5-dihydroxy-1,3-benzenedisulfonic acid disodium salt, Sigma), a O2–· scavenger, with dosage based on a number of previous studies that have used it effectively in this range without substantial toxicity (27, 34); 2) 50 µM Mn(III)tetrakis(1-methyl-4-pyridyl) porphyrin pentachloride (MnTMPyP, Oxis Research), a superoxide dismutase (SOD) mimic, with the dose chosen from cell culture studies to be extremely effective in protecting against oxidant stress with little toxicity (7, 31); and 3) 10 and 18 mM N-acetylcysteine (NAC, Sigma), a hydroxyl scavenger and precursor of glutathione, with dosage determined by previous studies of effective use in diaphragm skeletal muscle [10 mM (28) and 18 mM (34)]. All experiments were done in a pairwise manner. When necessary (e.g., NAC), the buffer solutions were titrated to pH 7.4 at 37°C when equilibrated to 5% CO2, before the tissues were exposed. Tiron (a mild acid) was equilibrated for ~15 min in well-bubbled buffer at 37°C to avoid excessive oxidation (yellow color), while providing sufficient time for the CO2-HCO3– buffer system to compensate for its mild acidification effect.

Antioxidant equilibration and hypoxia challenge.   The experimental protocol is illustrated in Fig. 1. Tissues were equilibrated in their respective treatment buffers for 30 min (Fig. 1, AOX Equilibration) while being stimulated with intermittent twitches (0.05 Hz). A new force-frequency maneuver was then performed at the end of the AOX incubation period (Fig. 1, Treatment FF). The strips were rested for 10 min and the preload was readjusted (Fig. 1, arrow). For the AOX experiments, each strip was put into one of four groups: 1) untreated time control in 95% O2 (control), 2) AOX treated in 95% O2 (control + AOX), 3) untreated hypoxic (hypoxia), and 4) AOX treated hypoxic (hypoxia + AOX). To test the response to hypoxia, the gas going to the hypoxia baths was changed to 95% N2-5% CO2 (hypoxia), and the buffers were replaced with buffer preequilibrated with 95% N2-5% CO2 at 37°C. The nonhypoxic baths (with and without AOX treatment) were changed to fresh buffer, preequilibrated with 95% O2-5% CO2 at 37°C (control). By use of a remote Clark PO2 detector (World Precision Instruments, Sarasota, FL), measurement of the PO2 within the hypoxic baths demonstrated values from 5–15 Torr near the center of the chamber after complete equilibration. The tissues remained in hypoxia or 95% O2 for 30 min. For the experimental series in which the extracellular Ca2+ was altered, the muscles were treated similarly, but there were no 95% O2 controls performed; the four muscles were exposed to four different Ca2+ conditions in hypoxia.

For the AOX treatments, two different stimulation paradigms were performed. In one paradigm (stimulated), muscles were intermittently stimulated with twitches (0.05 Hz) during hypoxic exposure to monitor contractile function during the time course of developing hypoxia, followed by a force-frequency test at the end of the hypoxic exposure. In the second paradigm (unstimulated), no twitch stimulations or force-frequency measurements were performed at any time during hypoxia. This latter group was necessary to eliminate the complexities of interpretation due to differing energy costs of contraction between AOX groups, i.e., because AOX-treated tissues generated more force in hypoxia, they presumably consumed more energy. All tissues used in the energetics study were rapidly freeze clamped across the center of the diaphragm muscle mass at the end of the hypoxic exposure period and then stored at –80°C. Our procedures allow us to freeze clamp the muscle within 1 s of dropping the surrounding tissue bath (24).

Extracellular Ca2+ experiments.   A separate series of experiments, designed to test the influence of extracellular Ca2+ on passive force during hypoxia, were performed in which [Ca2+] in the baths was manipulated (brackets denote concentration). Four diaphragm strips were mounted, and after the initial force-frequency, the baths were changed to control Ca2+ (2 mM), low Ca2+ (0 mM), 0 mM Ca2+ + 50 µM EGTA, or double Ca2+ (4 mM). To keep osmolarity approximately constant, changes in [NaCl] were used to compensate for changes in CaCl2.

Tissue extraction.   Frozen samples were weighed and ground in the presence of 0.6 N perchloric acid under liquid N2 by using liquid N2-cooled mortars and pestles. The N2 was allowed to evaporate, and the resulting powder was quickly placed in a vial containing additional ice-cold perchloric acid. Samples were vortexed and then centrifuged at 16,000 g for 1 min. One milliliter of supernatant was pipetted to a new tube, and the pellet was kept for protein analysis. To adjust the sample pH, 1 M K2CO3 was added to the supernatant and the sample was allowed to sit on ice for >30 min to precipitate KClO4 and then centrifuged. The final supernatant was pipetted into aliquots and stored at –80°C. All added volumes and weights were recorded for later analyses.

Metabolite analysis.   Pi was assayed by a modified colorimetric method by Fiske and Subbarow (17a). Briefly, 50 µl of thawed supernatant were added to 1.0 ml water and 1.0 ml assay buffer (0.5% ammonium molybdate, 2.0% ascorbic acid, and 2.8% concentrated sulfuric acid) and mixed, and the absorbance was read after 75 s at 820 nm on a spectrophotometer (Hewlett-Packard 8452A diode array). Concentrations were calculated on the basis of a standard curve. ATP and CrP are acid labile, so samples of known concentration were also measured to determine the portion of the signal attributable to their degradation. Timing of measurements was critical because of this degradation. Creatine, ATP, and CrP were assayed by the enzymatic methods of Bergmeyer (4). This procedure was described in detail in a previous manuscript (24). All assays were performed in duplicate and repeated if measurements varied by >15%. Final values were the means of the two closest values.

Protein assay.   Stored protein pellets were thawed and washed with acetone. Dried pellets were resuspended in 500 µl of 1 N NaOH and heated for 10 min at 95–100°C, then placed on ice or frozen at –20°C until assayed (standards were treated similarly). Total dissolved protein was determined by the Lowry method (32). Samples were read with a spectrophotometer at 750 nm and compared with a standard curve.

Statistical analysis.   Force measurements were always normalized to initial maximum force. Specific force, or stress, could not be determined in this study because tissues were immediately freeze clamped at the end of the hypoxic period, eliminating the ability to calculate the length and weight of the muscle. The assumption used was that before any treatments all tissues would exhibit equivalent baseline specific forces. Most analyses then used repeated-measures one-way or two-way ANOVA using JMP software (SAS Institute). Comparisons between means were tested, post-ANOVA, by use of contrast procedures. To evaluate differences between the amounts of contracture at the end of the hypoxic period, paired differences between matched controls were compared by using nonparametric matched-pairs statistics. Data points were expressed graphically as the average value ± SE.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Contracture during hypoxia.   Within 10–15 min of exposure to severe hypoxia, diaphragm strips exhibited gradual elevations in passive force, i.e., contracture, as illustrated in Fig. 2 (triangles). The data represent average values recorded from diaphragm strips stimulated at 0.05 Hz during exposure. In untreated hypoxic muscles (solid triangles), passive force generally increased 1 to 3 g during the 30-min exposure, or 1.5–2.0% of initial maximum tetanic force (150 Hz). The amount of contracture varied greatly between and within each experimental group but tended to vary less within animals. Therefore, paired nonparametric comparisons between individual strips from the same animals were necessary for statistical analyses, as shown in Fig. 3. Individual data points in this figure represent the final change in passive force after 30 min of hypoxic exposure. Pairs represent data from strips from the same animal treated with and without a given AOX. The experiments in which strips were stimulated with intermittent twitches during hypoxia are displayed on the left (squares), and experiments in which strips were unstimulated during hypoxia are on the right (triangles). In the strips stimulated during hypoxia, Tiron and MnTMPyP significantly reduced the level of contracture, whereas only Tiron significantly reduced contracture in the unstimulated strips. NAC (18 mM) had no consistent effect on contracture development (10 mM not shown). There were no significant effects of intermittent twitch stimulations during hypoxia on the overall extent of contracture in strips not treated with AOXs.



View larger version (26K):
[in this window]
[in a new window]
  		Fig. 2. Passive force as a percentage of initial maximum force (150 Hz) measured at 1-min intervals. Average values are passive force in tissues that were stimulated with or without exposure to hypoxia. Values are means ± SE; n = 6 for each AOX. Tiron, 4,5-dihydroxy-1,3-benzenedisulfonic acid; MnTMPyP, Mn(III)tetrakis(1-methyl-4-pyridyl) porphyrin pentachloride; NAC, N-acetylcysteine.

 


View larger version (16K):
[in this window]
[in a new window]
  		Fig. 3. Changes in passive force from initial during hypoxic exposure, expressed as a percentage of initial maximum force (150 Hz). Each data point is from individual experiments and represents the total change in passive force during hypoxia exposure (squares, stimulated; triangles, unstimulated during hypoxia). The lines match paired tissues in control and AOX-treated tissues from the same animal.

 
Changes in extracellular [Ca2+] had no effect on passive force or on the force-frequency relationship after 30 min of incubation, before hypoxia (data not shown). However, during hypoxia, contracture was essentially abolished by removal of extracellular Ca2+, as shown in Fig. 4. We could identify no significant effect of doubling extracellular [Ca2+] on the extent of contracture.



View larger version (18K):
[in this window]
[in a new window]
  		Fig. 4. Effect of extracellular Ca2+ concentration on passive force during hypoxia. Values are means ± SE; n = 6 for 0 mM, control, and 4 mM; n = 5 for 0 mM + EGTA. *P < 0.05 compared with control Ca2+.

 
Contractile function in hypoxia.   Figure 5 illustrates the average peak twitch forces measured at the end of the 30 min of antioxidant incubation (hatched bars, n = 12, all tissues with and without AOX before hypoxia) and at the end of the 30 min of hypoxia for each stimulated treatment group (solid bars, n = 6). Results are expressed as a percent of the initial maximal force. Note that, compared with matched 95% O2 controls, Tiron, NAC, and MnTMPyP decreased peak twitch force, as expected (12, 37). Interestingly, after 30 min of hypoxia, only MnTMPyP significantly increased twitch force compared with matched, untreated hypoxia tissues.



View larger version (36K):
[in this window]
[in a new window]
  		Fig. 5. Effects of antioxidants on twitch force before and after 30 min of hypoxia as a percentage of initial maximum tetanic force. Hatched bars, data after 30-min incubation in normoxia (n = 12). Solid bars, data after 30 min of hypoxia (n = 6). Values are means ± SE. *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001. Statistical differences are from post-ANOVA contrasts for individual AOX treatments vs. non-AOX treated matched controls.

 
As shown by the solid symbols for untreated strips in Fig. 6, the force-frequency curves were shifted down and to the right after 30 min of exposure to hypoxia. The relative degree of hypoxic-induced depression of force is more evident at low frequencies (e.g., 20 Hz force was reduced ~85% of matched control values) compared with high frequencies (150-Hz force was reduced only ~50–60% of matched control values). Antioxidant treatment in strips not exposed to hypoxia (open circles) resulted in reductions in force at nearly all frequencies, suggesting that, in this setting, high levels of AOXs may have a long-term (>1 h exposure) negative impact on overall contractile function. This did not reach statistical significance for NAC-treated tissues, except at low frequencies.



View larger version (20K):
[in this window]
[in a new window]
  		Fig. 6. Force-frequency relationships measured at the end of hypoxia as a percentage of initial maximum tetanic force. Tissues studied at 95% O2 are time-matched controls, and each graph represents 1 antioxidant [top, Tiron; middle, superoxide dismutase (SOD) mimic; bottom, NAC]. Values are means ± SE; n = 6 for each AOX. *P ≤ 0.05, **P ≤ 0.01.

 
Treatment with the two O2–· scavengers, Tiron or MnTMPyP, resulted in significant preservation of force at all but the lowest stimulation frequencies for tissues exposed to hypoxia. NAC (18 mM) had no influence on the force-frequency relationship after 30 min of hypoxia exposure. When NAC studies were repeated at 10 mM NAC, no significant effects were observed (not shown).

High-energy phosphate measurements in hypoxia.   High-energy phosphates and Pi were measured in fresh frozen samples, strips that were stimulated during hypoxia, and strips that were not stimulated during hypoxia. Results are illustrated in Fig. 7. Data for NAC experiments are not shown for simplicity, because there was no protection of contractile function and they could not be used to test the primary hypothesis of the study. However, measurements were performed for NAC-treated muscles (10 mM) contracting during hypoxia. No significant effects of NAC treatment on ATP, CrP, or Pi were observed (not shown). In Fig. 7A, the [ATP] [~0.05 µmol/mg protein, corresponds to ~3.5 µmol/g wet wt (24)] of fresh frozen diaphragm (open bars) was similar to control diaphragm strips incubated in the tissue baths. In hypoxic muscle strips that were not stimulated during hypoxia there was a small trend for [ATP] to decrease, but this did not reach statistical significance. However, strips that were electrically stimulated to twitch and to generate a force-frequency relationship during hypoxia showed significant (~20–30%) reductions in [ATP] compared with matched, stimulated controls. Only Tiron treatment during hypoxia resulted in no significant reduction in [ATP] compared with matched controls in these tissues. When the values for hypoxia and hypoxia + Tiron were tested against each other, however, there were no differences in these groups. In assessing the overall response of all tissue groups, the lack of effect by hypoxia in Tiron-treated strips appears to be similar to the results for other antioxidant treatments.



View larger version (32K):
[in this window]
[in a new window]
  		Fig. 7. High-energy phosphate measurements in hypoxia and time-matched controls, normalized to micrograms protein for each group. Hatched bars, tissues stimulated during hypoxia. Solid bars, no stimulation during hypoxia. *P ≤ 0.05. Values are mean ± SE; n = 6 for Tiron and MnTMPyP, n = 4 for fresh frozen.

 
CrP concentrations (Fig. 7B) in fresh frozen tissue were roughly the same as in all control tissues, averaging ~0.15 µmol/mg protein, corresponding to ~11.6 µmol/g wet wt (24). Hypoxia exposure drastically reduced CrP to ~10% of normoxic values. There were no significant effects of any AOX treatment compared with nontreated control tissues, although in unstimulated tissues CrP tended to remain higher at the end of hypoxia compared with the stimulated strips.

As shown in Fig. 7C, Pi increased significantly during hypoxia, approaching values corresponding to the reduction in CrP. There were no significant effects of antioxidant treatment on Pi in any of the tissues studied.


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
The results demonstrate that diaphragm muscles treated with O2–· scavengers during hypoxic exposure can attain ~30% higher maximum tetanic forces compared with untreated tissues, with no apparent additional reserves of high-energy phosphates. Therefore, the primary hypothesis of this study was disproved. Antioxidants also inhibited the development of hypoxia-induced contracture. Because contracture was shown to be dependent on extracellular Ca2+, implying a Ca2+-dependent mechanism, the results suggest that O2–· scavengers may impact hypoxia-induced alterations in Ca2+ regulation.

Comparison with previous studies and critique of the approach.   The results for the effects of AOXs on contractile function confirm our preceding work with Tiron under similar conditions (34). However, in contrast to our previous work, NAC had little or no influence on tetanic force production in hypoxia. A number of exploratory experiments were performed in an attempt to resolve this discrepancy within our laboratory (data not shown), namely 1) several different formulations and concentrations of NAC were compared, 2) differences in the time of NAC incubation were evaluated (30 min–2 h), and 3) the original fabricated field-stimulating electrodes were tested against currently employed commercial electrodes because of the potential effects of metal chemistry in buffer systems. None of these experiments reconciled the differences in results. We conclude that some unknown variable, e.g., the source of animals, diet, the source of distilled water, or metal-contaminating buffer salts had changed between the sets of experiments. NAC did have the expected influence on twitch force, before hypoxia, as shown in Fig. 5, confirming the previously described pharmacological effects of this agent on skeletal muscle (12, 28, 34).

The present results differ from the recent work of Heunks et al. (23), who found that AOXs, Tiron, and NAC depressed maximum force development during hypoxia compared with hypoxia alone. Differences in the experimental protocols between the two studies suggest that the level of hypoxia was much more severe in the present study. First, bath temperature was 26°C in Heunks et al. vs. 37°C in the present study. Therefore, metabolic rate would be higher and dissolved O2 lower at 37°C, predictably making the tissues in this experiment more hypoxic in the central core of the muscle bundle. Second, the PO2 was ~7.5 kPa (~56 Torr) for 60 min vs. ~1–2 kPa (~5.0–15.0 Torr) for 30 min in this study. Third, the average animal size in the Heunks study was ~300 g vs. ~400 g in the present study, which could have resulted in thicker diaphragm tissues and a greater core hypoxia in the present study. Therefore, it is not surprising that the maximum tetanic force we observed after 30 min of hypoxia was ~50% of baseline compared with ~90% of baseline in the study of Heunks et al. after 60 min of hypoxia.

As pointed out in a number of recent publications (15, 16), the redox-dependent responses to hypoxia in skeletal muscle vary widely within a relatively narrow range of PO2 exposure. A rough estimate of the PO2 as a function of tissue depth in isolated skeletal muscle is provided from work by Eu et al. (15) in resting mouse extensor digitorum longus muscle, which is ~1 mm in thickness, compared with 0.4–0.6 mm in rat diaphragm. When bath PO2 is 1% O2, the PO2 at the core of the extensor digitorum longus was ~3–4 Torr; at a 20% O2 core PO2 was ~40 Torr; at 95% O2 core PO2 was 387 Torr. These measurements are extremely difficult to make accurately and should be viewed with some caution; nevertheless they suggest that, in a hypoxic tissue bath, PO2 reductions in the muscle core can be substantial. Normal PO2 in perfused skeletal muscle is believed to be ~10 Torr at rest and 3–5 Torr during intense exercise (38), not reaching a "critical" cell PO2 where respiration is limited until ~1.5 Torr (39). The present study likely resulted in relatively severe hypoxia in the muscle core with more moderate hypoxia in the study by Heunks et al. (23). We speculate that different response mechanisms predominate at different levels of severity of hypoxia and therefore it is difficult but interesting to compare the results from the two laboratories.

Influences of specific AOXs.   It is of some interest that the two O2–· scavengers had such potent impacts on contractile function in this setting whereas NAC, a more commonly used antioxidant or reducing agent, had only small effects. O2–· is not considered to be extremely reactive as an oxidant and behaves as a reductant in many microenvironments. However, some targets, such as isoforms of the sarcoplasmic reticulum (SR) Ca2+-ATPase, appear to be specifically sensitive to O2–· oxidation (3) whereas other targets may be uniquely sensitive to peroxynitrite, resulting from reactions that require O2–·.

It is instructive to compare the relative scavenging abilities of Tiron and MnTMPyP to the endogenous enzyme, SOD. Endogenous SOD activity in rat diaphragm has been estimated to be ~3.5 U/mg (30). By taking the approach of Gardner et al. (19), assuming a soluble protein concentration of 80 mg/ml in diaphragm (this study), the endogenous SOD concentration can be estimated at ~2.5 µM. Because SOD has a reaction rate with O2–· of ~109 M–1·s–1 it would scavenge at a rate of 2.5 x 103 s–1. By contrast, Tiron scavenges O2–· with a rate constant of ~5 x 108 M–1·s–1 (21), and, assuming a 10 mM intracellular concentration, it would scavenge O2–· at 5 x 106 s–1, a 2,000 times greater rate than endogenous SOD. Similarly, assuming that MnTMPyP is kept in the reduced state [Mn(II)·TMPyP] by cellular reductants, the Mn(II)TMPyP has a rate constant of 4 x 109 M–1·s–1 for oxidation by O2–· (17) and intracellular concentrations of 50 µM would result in a net reaction rate of 2 x 105 s–1, 80-fold higher SOD activity than endogenous SOD. It is highly likely that intracellular concentrations of these applied scavengers are considerably less than their extracellular concentrations and that endogenous SOD is more effective than suggested by these calculations because it is strategically located near O2–·-generating sites such as the mitochondria. Nevertheless, the calculations demonstrate that the scavengers used in this study were extremely potent at the concentrations used. The lack of effect of NAC on the response may simply reflect its comparatively reduced scavenging ability. It is known primarily as a hydroxyl scavenger and precursor for glutathione synthesis. It is not known to have significant scavenging effects on O2–· and only slowly reacts with H2O2 (2).

It is also important to note that neither of the O2–· scavengers is particularly specific. Tiron is an effective metal scavenger (27) and MnTMPyP acts as both a catalase (7) and a peroxynitrite scavenger (18) as well. We infer that it is likely that these AOXs affected a variety of reactive O2 and reactive nitrogen species and therefore the specificity of O2–· in this physiological response to hypoxia should be viewed with some caution.

Potential mechanisms of antioxidant inhibition of contracture.   The results showed that contracture could be eliminated by removal of extracellular Ca2+, suggesting that the phenomenon is related to hypoxia-induced alterations in cell Ca2+ homeostasis. Several known mechanisms could account for the influence of AOXs on Ca2+ homeostasis. First, movement of Ca2+ into the cytosol would be expected in response to membrane depolarization, from either the SR or possibly the extracellular fluid. Membrane depolarization is known to occur in rat diaphragm during hypoxia (14). Second, oxidants could influence K+ channel behavior in hypoxia and thus affect Ca2+ influx indirectly. In general, the membrane resists depolarization by outward K+ leakage and recruitment of new K+ channels. One of the molecular mechanisms believed to be responsible for O2 sensing is the influence of local redox state on K+ channel behavior (33), although little is known regarding these influences in skeletal muscle. Third, the SR Ca2+ release channel in skeletal muscle (RyR1) is sensitive to the local redox environment. The open probability of the channel is highly dependent on complex interactions between oxidants, O2, and NO (1, 15, 16, 40). This effect has been described as a form of O2 sensing by RyR1 (15, 16). Fourth, the SR Ca2+-ATPase is responsible for clearance of cytosolic Ca2+ in skeletal muscle, and it is dependent on the presence of an effective creatine kinase system and available CrP (8, 13). Because CrP dropped below 10% of normal during hypoxia, a value shown to inhibit SR Ca2+-ATPase activity in skinned skeletal muscle (13), it is likely that low Ca2+-ATPase pump activity contributed to the accumulation of cytosolic Ca2+. Nearly all isoforms of the Ca2+-ATPase are redox sensitive, with oxidants causing a reduction in function (3, 41). Recent evidence suggests that at least some isoforms of Ca2+-ATPase in skeletal muscle are uniquely inhibited by O2–· (3). Fifth, cell swelling may also contribute to the elevation in passive force during hypoxia. Presumably, as muscle fibers swell, stresses on the elastic elements of the cytoskeleton and membrane could exert passive forces along the long axis of the fiber, contributing to contracture. Swelling occurs in the skeletal muscle in the setting of hypoxia due to the additional osmotic pressure caused by the splitting of CrP to Pi and Cr and the degradation of glycogen to glucose and eventually lactate (29). Interestingly, cell swelling causes an increase in intracellular Ca2+ and subsequent reactive O2 formation via a phospholipase A2-dependent pathway (36).

Mechanisms of tetanic force preservation in hypoxia.   Because AOXs did not preserve high-energy phosphates, other mechanisms for preservation of contractile function must be considered. One possible explanation is that AOXs preserved tetanic force in hypoxia by allowing the muscles to contract more economically. Economy is defined as tetanic force developed per rate of ATP hydrolysis. Although the specific rate of ATP hydrolysis was not measured during contraction, the data suggest that there was little or no apparent additional energetic cost to contract at higher forces with AOX treatment because there was no additional decrement in [ATP] and [CrP]. This conclusion is somewhat limited by the inherent inaccuracies involved in the measurement of high-energy phosphates in frozen samples, as discussed previously (24), and because of the fact that the actual rate of hydrolysis was not measured. Furthermore, we cannot rule out the possibility that the rate of ATP turnover was elevated in AOX-treated tissues. It is possible that at specific levels of CrP, ATP, and ADP during hypoxia, the metabolic network is able to maintain better coupling of energy utilization and energy production.

One potential mechanism that could account for improved economy of contraction in hypoxia is a more homogeneous distribution of forces along the fiber length. For example, if certain regions of a muscle, in series along a fiber, are poorly activated during hypoxia compared with other better activated areas, the activated areas will shorten disproportionately and stretch the noncontracting portions of the fibers, thus reducing the net force transmission from tendon to tendon. In hypoxic muscle, it is likely that some regions of a fiber are more vulnerable to energy depletion. Recent data suggest that within sarcomeric regions there is a compartmentalization of energy metabolic systems, at least in cardiac muscle (35), and this could vary with depth inside the tissue and fiber. Alternatively, some other less defined redox-sensitive mechanism that influences the efficiency of cross-bridge interaction could be responsible for the effects of antioxidant treatment. For example, the downward shift in the force-velocity relationship (a characteristic of slower more efficient muscles) was observed with Tiron treatment in more moderate hypoxia (23), suggesting that such a mechanism is possible.

Some of the same potential influences of AOXs on Ca2+ regulation that were discussed in the previous section regarding contracture may also apply to the development of tetanic force in hypoxia. For example, oxidative inhibition of Ca2+-ATPase (3, 41) would presumably lead to reductions in SR Ca2+ stores in hypoxia and therefore reductions in maximum Ca2+ release and maximum force development during contraction. Similarly, oxidation of ion channel activity may limit propagation of the action potential down T tubules, resulting in poor distribution of excitation-contraction coupling in some areas of the muscle. Antioxidants could work in hypoxia by blocking these effects, and the result would be that the muscle would contract in a more homogenous way and presumably do so more economically.

Physiological implications.   It is simplistic to conclude that hypoxia represents a form of "oxidative stress" in which AOXs are protective. It is probably more reasonable to consider the low level of oxidants produced in hypoxia as one of many signaling events that are necessary to set up some secondary protective system. For example, is it a good thing to be able to generate more force in a severe hypoxic environment when ATP and CrP are depleted as were observed in the presence of AOXs in this study? Contractions during hypoxia have been shown to cause damage to muscle proteins (9). From the data, it is clear that simply performing a force-frequency curve in hypoxia results in significant decrements in [ATP], whereas [ATP] is reasonably protected in the rested hypoxic muscle. Much like mechanisms of fatigue, muscles are likely to be programmed to protect themselves from overexertion and injury in such environments. For example, in a previous study in heart muscle by Klawitter et al. (25), Tiron was particularly effective in preserving high-energy phosphates in the heart when administered during 20 min of ischemia. However, it resulted in large decrements in energy status in the reperfusion phase, suggesting that oxidants produced during ischemia represent a desired protective mechanism in the heart, possibly providing a type of preconditioning stimulus. Similar mechanisms may be present in skeletal muscle but possibly working through pathways that differ from those in the heart.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
This research was supported by National Heart, Lung, and Blood Institute Grant HL-53333.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
REFERENCES
 
Thanks go to Valery Khramtsov for discussions regarding biochemical aspects of the manuscript.

Present address of P. Klawitter: Dept. of Emergency Medicine, Upstate Medical Univ., 750 E. Adams St., Syracuse, NY 13210.


   FOOTNOTES
 

Address for reprint requests and other correspondence: T. Clanton, The Ohio State Univ., Dept. of Internal Medicine, Pulmonary, Critical Care and Sleep Medicine, Rm. 201, Dorothy M. Davis Heart & Lung Research Institute, 473 W 12th, Columbus, OH 43210 (E-mail: clanton.1{at}osu.edu)

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.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 

  1. Abramson J and Salama G. Sulfhydryl oxidation and Ca2+ release from the sarcoplasmic reticulum. Mol Cell Biochem 82: 81–84, 1988.[CrossRef][ISI][Medline]
  2. Arumona OI, Halliwell B, Hoey BM, and Butler J. The antioxidant action of N-acetylcysteine: its reaction with hydrogen peroxide, hydroxyl radical, superoxide and hypochlorous acid. Free Radic Biol Med 6: 593–597, 1989.[CrossRef][ISI][Medline]
  3. Barnes KA, Samson SE, and Grover AK. Sarco/endoplasmic reticulum Ca2+-pump isoform SERCA3a is more resistant to superoxide damage than SERCA2b. Mol Cell Biochem 203: 17–21, 2000.[CrossRef][ISI][Medline]
  4. Bergmeyer HU (Editor). Methods of Enyzmatic Analysis (3rd ed.). Deerfield Beach, FL: Verlag Chemie, 1985.
  5. Blair AS, Hajduch E, Litherland GJ, and Hundal HS. Regulation of glucose transport and glycogen synthesis in L6 muscle cells during oxidative stress. Evidence for cross-talk between the insulin and SAPK2/p38 mitogen-activated protein kinase signaling pathways. J Biol Chem 274: 36293–36299, 1999.[Abstract/Free Full Text]
  6. Day BJ, Fridovich I, and Crapo JD. Manganic porphyrins possess catalase activity and protect endothelial cells against hydrogen peroxide-mediated injury. Arch Biochem Biophys 347: 256–262, 1997.[CrossRef][ISI][Medline]
  7. De Groof AJ, Fransen JA, Errington RJ, Willems PH, Wieringa B, and Koopman WJ. The creatine kinase system is essential for optimal refill of the sarcoplasmic reticulum Ca2+ store in skeletal muscle. J Biol Chem 277: 5275–5284, 2002.[Abstract/Free Full Text]
  8. De Paula Brotto M, van Leyen SA, Brotto LS, Jin JP, Nosek CM, and Nosek TM. Hypoxia/fatigue-induced degradation of troponin I and troponin C: new insights into physiologic muscle fatigue. Pflügers Arch 442: 738–744, 2001.[CrossRef][ISI][Medline]
  9. Depre C, Rider MH, and Hue L. Mechanisms of control of heart glycolysis. Eur J Biochem 258: 276–290, 1998.
  10. Depre C, Vanoverschelde JL, Goudemant JF, Mottet I, and Hue L. Protection against ischemic injury by nonvasoactive concentrations of nitric oxide synthase inhibitors in perfused rabbit heart. Circulation 92: 1911–1918, 1995.[Abstract/Free Full Text]
  11. Diaz PT, Brownstein E, and Clanton TL. Effects of N-acetylcysteine on in vitro diaphragm function are temperature dependent. J Appl Physiol 77: 2434–2439, 1994.[Abstract/Free Full Text]
  12. Duke AM and Steele DS. Effects of creatine phosphate on Ca2+ regulation by the sarcoplasmic reticulum in mechanically skinned rat skeletal muscle fibers. J Physiol 517: 447–458, 1999.[Abstract/Free Full Text]
  13. Esau SA. Role of adenosine in the depolarization of hypoxic hamster diaphragm membrane in vitro. Am J Respir Crit Care Med 149: 910–914, 1994.[Abstract]
  14. Eu JP, Hare JM, Hess DT, Skaf M, Sun J, Cardenas-Navina I, Sun QA, Dewhirst M, Meissner G, and Stamler JS. Concerted regulation of skeletal muscle contractility by oxygen tension and endogenous nitric oxide. Proc Natl Acad Sci USA 100: 15229–15234, 2003.[Abstract/Free Full Text]
  15. Eu JP, Sun J, Xu L, Stamler JS, and Meissner G. The skeletal muscle calcium release channel: coupled O2 sensor and NO signaling functions. Cell 102: 499–509, 2000.[CrossRef][ISI][Medline]
  16. Faulkner KM, Liochev SI, and Fridovich I. Stable Mn(III) porphyrins mimic superoxide dismutase in vitro and substitute for it in vivo. J Biol Chem 269: 23471–23476, 1994.[Abstract/Free Full Text]
  17. Fiske CH and Subbarow Y. The colorimetric determination of phosphorus. J Biol Chem 66: 375–400, 1925.[Free Full Text]
  18. Ferrer-Sueta G, Batinic-Haberle I, Spasojevic I, Fridovich I, and Radi R. Catalytic scavenging of peroxynitrite by isomeric Mn(III) N-methylpyridylporphyrins in the presence of reductants. Chem Res Toxicol 12: 442–449, 1999.[CrossRef][ISI][Medline]
  19. Gardner PR, Nguyen DD, and White CW. Superoxide scavenging by Mn(II/III) tetrakis (1-methyl-4-pyridyl) porphyrin in mammalian cells. Arch Biochem Biophys 325: 20–28, 1996.[CrossRef][ISI][Medline]
  20. Godt RE and Nosek TM. Changes in intracellular milieu with fatigue or hypoxia depress contraction of skinned rabbit skeletal and cardiac muscle. J Physiol 412: 155–180, 1989.[Abstract/Free Full Text]
  21. Greenstock CL and Miller RW. The oxidation of Tiron by superoxide anion: kinetics of the reaction in aqueous solution and in chloroplasts. Biochim Biophys Acta 396: 11–16, 1975.[Medline]
  22. Henry PD, Schuchleib R, Davis J, Weiss ES, and Sobel BE. Myocardial contracture and accumulation of mitochondrial calcium in ischemic rabbit heart. Am J Physiol Heart Circ Physiol 233: H677–H684, 1977.[Abstract/Free Full Text]
  23. Heunks LM, Machiels HA, de Abreu R, Zhu XP, van der Heijden HF, and Dekhuijzen PN. Free radicals in hypoxic rat diaphragm contractility: no role for xanthine oxidase. Am J Physiol Lung Cell Mol Physiol 281: L1402–L1412, 2001.[Abstract/Free Full Text]
  24. Klawitter PF and Clanton TL. Tension-time index, fatigue, and energetics in isolated diaphragm: a new experimental model. J Appl Physiol 96: 89–95, 2004.[Abstract/Free Full Text]
  25. Klawitter PF, Murray HN, Clanton TL, and Angelos MG. Reactive oxygen species generated during myocardial ischemia enable energetic recovery during reperfusion. Am J Physiol Heart Circ Physiol 283: H1656–H1661, 2002.[Abstract/Free Full Text]
  26. Kowaltowski AJ and Vercesi AE. Mitochondrial damage induced by conditions of oxidative stress. Free Radic Biol Med 26: 463–471, 1999.[CrossRef][ISI][Medline]
  27. Krishna CM, Liebmann JE, Kaufman D, DeGraff W, Hahn SM, McMurry T, Mitchell JB, and Russo A. The catecholic metal sequestering agent 1,2-dihydroxybenzene-3,5-disulfonate confers protection against oxidative cell damage. Arch Biochem Biophys 294: 98–106, 1992.[CrossRef][ISI][Medline]
  28. Kwali FA and Reid MB. N-acetylcysteine depressed contractile function and inhibits fatigue of diaphragm in vitro. J Appl Physiol 77: 317–324, 1994.[Abstract/Free Full Text]
  29. Lang F, Busch GL, Ritter M, Völkl H, Waldegger S, Gulbins E, and Häussinger D. Functional significance of cell volume regulatory mechanisms. Physiol Rev 78: 247–306, 1998.[Abstract/Free Full Text]
  30. Lawler JM, Powers SK, Dijk HV, Visser T, Kordus MJ, and Ji LL. Metabolic and antioxidant enzyme activities in the diaphragm: effects of acute exercise. Respir Physiol 96: 139–149, 1994.[CrossRef][ISI][Medline]
  31. Lee JH and Park JW. A manganese porphyrin complex is a novel radiation protector. Free Radic Biol Med 37: 272–283, 2004.[CrossRef][Medline]
  32. Lowry OH, Rosebrough NJ, Farr AL, and Randall RJ. Protein measurement with Folin phenol reagent. J Biol Chem 193: 265–275, 1951.[Free Full Text]
  33. López-Barneo J, del Toro R, Levitsky KL, Chiara MD, and Ortega-Sáenz P. Regulation of oxygen sensing by ion channels. J Appl Physiol 96: 1187–1195, 2004.[Abstract/Free Full Text]
  34. Mohanraj P, Merola AJ, Wright VP, and Clanton TL. Antioxidants protect rat diaphragmatic muscle function under hypoxic conditions. J Appl Physiol 84: 1960–1966, 1998.[Abstract/Free Full Text]
  35. Ovádi J and Saks V. On the origin of intracellular compartmentation and organized metabolic systems. Mol Cell Biochem 256/257: 5–12, 2004.[CrossRef]
  36. Ørtenblad N, Young JF, Oksbjerg N, Nielsen JH, and Lambert IH. Reactive oxygen species are important mediators of taurine release from skeletal muscle cells. Am J Physiol Cell Physiol 284: C1362–C1373, 2003.[Abstract/Free Full Text]
  37. Reid MB, Khawli F, and Moody MR. Reactive oxygen in skeletal muscle III. Contractility of unfatigued muscle. J Appl Physiol 75: 1081–1087, 1993.[Abstract/Free Full Text]
  38. Richardson RS, Noyszewski EA, Saltin B, and Gonzalez-Alonso J. Effect of mild carboxy-hemoglobin on exercising skeletal muscle: intravascular and intracellular evidence. Am J Physiol Regul Integr Comp Physiol 283: R1131–R1139, 2002.[Abstract/Free Full Text]
  39. Richmond KN, Burnite S, and Lynch RM. Oxygen sensitivity of mitochondrial metabolic state in isolated skeletal and cardiac myocytes. Am J Physiol Cell Physiol 273: C1613–C1622, 1997.[Abstract/Free Full Text]
  40. Salama G, Menshikova EV, and Abramson JJ. Molecular interaction between nitric oxide and ryanodine receptors of skeletal and cardiac sarcoplasmic reticulum. Antioxid Redox Signal 2: 5–16, 2000.[Medline]
  41. Suzuki YJ and Ford GD. Inhibition of Ca2+-ATPase of vascular smooth muscle sarcoplasmic reticulum by reactive oxygen intermediates. Am J Physiol Heart Circ Physiol 261: H568–H574, 1991.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Exp PhysiolHome page
A. Dutta, K. Ray, V. K. Singh, P. Vats, S. N. Singh, and S. B. Singh
L-carnitine supplementation attenuates intermittent hypoxia-induced oxidative stress and delays muscle fatigue in rats
Exp Physiol, October 1, 2008; 93(10): 1139 - 1146.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
T. L. Clanton
Hypoxia-induced reactive oxygen species formation in skeletal muscle
J Appl Physiol, June 1, 2007; 102(6): 2379 - 2388.
[Abstract] [Full Text] [PDF]

Home page
 J. Physiol.Home page
M. E. Sandstrom, S.-J. Zhang, J. Bruton, J. P. Silva, M. B. Reid, H. Westerblad, and A. Katz
Role of reactive oxygen species in contraction-mediated glucose transport in mouse skeletal muscle
J. Physiol., August 15, 2006; 575(1): 251 - 262.
[Abstract] [Full Text] [PDF]

Home page
 Exp PhysiolHome page
J. D. Pierce, C. Goodyear-Bruch, S. Hall, and R. L. Clancy
Effect of dopamine on rat diaphragm apoptosis and muscle performance
Exp Physiol, July 1, 2006; 91(4): 731 - 740.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
M. C. Gong, S. Arbogast, Z. Guo, J. Mathenia, W. Su, and M. B. Reid
Calcium-independent phospholipase A2 modulates cytosolic oxidant activity and contractile function in murine skeletal muscle cells
J Appl Physiol, February 1, 2006; 100(2): 399 - 405.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Lung Cell. Mol. Physiol.Home page
C. A. C. Ottenheijm, L. M. A. Heunks, M. C. P. Geraedts, and P. N. R. Dekhuijzen
Hypoxia-induced skeletal muscle fiber dysfunction: role for reactive nitrogen species
Am J Physiol Lung Cell Mol Physiol, January 1, 2006; 290(1): L127 - L135.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
L. Zuo and T. L. Clanton
Reactive oxygen species formation in the transition to hypoxia in skeletal muscle
Am J Physiol Cell Physiol, July 1, 2005; 289(1): C207 - C216.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
98/5/1753    most recent
01022.2004v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (9)
Right arrow Citing Articles via Google Scholar
Google Scholar