| | Ischemia/reperfusion injury of skeletal muscle: Plasma taurine as a measure of tissue damage☆☆☆★Accepted 26 June 2002. Abstract Background. Cell membrane rupture by oxygen-derived free radicals is a systematic feature of ischemia/reperfusion (I/R) injury. High taurine concentration gradients in skeletal muscle prompted us to evaluate whether plasma taurine levels (pTau) are a useful marker of I/R injury after different periods of ischemia. Methods. Rabbits were randomly assigned to either 1 or 2.5 hours of hind-limb ischemia followed by 2 hours of reperfusion (groups IR1 [n = 12] and IR2.5 [n = 13], respectively). Corresponding sham groups (SHAM1 [n = 8] and SHAM2.5 [n = 9]) were used as controls. Analyzed parameters included histomorphometry and electron microscopy of skeletal muscle biopsies, pTau, and plasma level of malondialdehyde. Skeletal muscle function was assessed 3 weeks after I/R injury. Results. No significant morphologic changes were detectable at the end of ischemia. After reperfusion, mild interstitial edema with intact muscle cell membranes developed in IR1 group; pTau was not increased. IR2.5 group, by contrast, showed severe interstitial edema formation (interfiber area increased by 112%, P < .005), microvascular constriction (microvessel area decreased by 33%, P < .0005), and damage to the muscle cell membranes that was confirmed by the increased plasma malondialdehyde. pTau was higher than in the SHAM2.5 group (P < .0005). Pronounced cell damage in IR2.5 group resulted in impaired muscle function (maximal tetanic tension was reduced 2 times, P < .005) but not in IR1 group. Conclusion. Skeletal muscle tolerates 1 h/2 h but not 2.5 h/2 h of I/R, the latter resulting in interstitial edema formation, microvascular constriction, and a late muscle dysfunction. Cell membrane rupture through stimulated lipid peroxidation promotes leakage of intracellular taurine, leading to increased pTau after reperfusion and may be considered as prognostically unfavorable in terms of organ function reversibility. In the rabbit model, pTau seems to be a sensitive marker of I/R injury to skeletal muscle. (Surgery 2003;133:91-100.)
Re-establishment of blood flow after ischemia adds paradoxically to the damage done by prolonged ischemia.1 This phenomenon is called ischemia/reperfusion (I/R) injury, which is characterized by perfusion disorders ("no reflow") and interstitial edema formation as a result of capillary constriction and increased permeability.2
Reperfusion of ischemic tissue results in the generation of superoxide anion2, 3, 4 and other extremely reactive oxygen-derived free radicals. Oxygen-derived free radicals cause lipid peroxidation5 and, consequently, disruption of the sarcolemmal integrity impairing the transport functions of cell membranes.6 The pathways of superoxide anion production include deranged constitutive nitric oxide synthase,7 hypoxanthine-xanthine oxidase pathway,8 activation of polymorphonuclear leukocytes,9 mitochondrial electron transport, catecholamine oxidation, and prostaglandin metabolism.1
Taurine (2-aminoethane sulfonic acid) is a ubiquitous sulfur-containing amino acid present in various tissues. The bulk of taurine is in the musculature, even though its highest concentrations are in the heart and brain.10 Taurine involves unusually high concentration gradients across cell membranes,11 such as 300:1 in human skeletal muscle.12 This high gradient is maintained by the lipophobic properties of its β-amino acid, which effectively prevent taurine from diffusing through lipophilic cell membranes.11 The entry of taurine into the cells is by means of a specific carrier-mediated, sodium cation (Na+) dependent transport system stimulated by chloride ion (Cl−) (Na+/Cl− coupled transport).11
The leakage of taurine and increased taurine blood and plasma concentrations (pTau) were described in patients undergoing coronary bypass operation,13 after myocardial infarction,14 or myocardial ischemia.15 However, data about the duration of ischemia leading to taurine leakage are lacking.
We hypothesized that only a certain degree of I/R injury that causes sarcolemmal disruption of muscle fibers and compromises active cellular transport system as a result of depletion of cellular energy stores could lead intracellular taurine (iTau) to leak into the interstitial space and, as a result, raise pTau. Therefore, we performed the current study to investigate I/R injury of skeletal muscle after different ischemic intervals and to assess pTau as a possible marker of cell membrane rupture. The pTau changes were compared with other markers of tissue injury (creatine kinase [CK], lactate dehydrogenase [LDH]). Moreover, long-term functional sequelae of different periods of ischemia on the skeletal muscle were investigated to correlate increased pTau during I/R injury with reversibility of impaired muscle function.
Material and methods  Surgical procedure The study was carried out on adult male New Zealand white rabbits (Charles River GmbH, Sulzfeld, Germany) weighing 2.7 to 3.5 kg. The experiments were approved by the local ethics committee and were in conformity with the Declaration of Helsinki (Guiding Principles for Research Involving Human Beings and Animals) and the Guiding Principles in the Care and Use of Animals (approved by the Council of the American Physiological Society). The rabbits were allowed to move freely in their cages and had free access to food (Altromin 2120 standard diet pellets (Marek, Vienna, Austria) and water. Food was withdrawn 24 hours before operation. The animals were randomly allocated to 4 experimental groups: the IR1 (n = 12) and IR2.5 (n = 13) groups were subjected to hind-limb I/R comprising either 1 or 2.5 hours of ischemia followed by 2 hours of reperfusion. The SHAM1 (n = 8) and SHAM2.5 (n = 9) control groups were subjected to sham operations covering the same periods of time with no I/R injury involved. The rabbit model of hind-limb I/R injury used in this study was previously described in detail.7 In brief, intravenous narcosis was attained using a mixture of 15 mg/kg ketamine hydrochloride (Ketalar, Parke-Davis GmbH, Berlin, Germany) and 1.5 mg/kg xylazine hydrochloride (Rompun, Bayer, Leverkusen, Germany). Next, the animals were intubated by a tracheotomy. Lung ventilation was maintained with nitrous oxide/oxygen inhalation (N2/O2) (FI O2 [fraction of inspired oxygen] = 0.35) and isoflurane (Forane; Abbott, Vienna, Austria) 1 to 2 vol percent at a tidal volume of 15 to 20 mL/kg and a rate of 30 to 35 cycles/min. Systemic blood pressure was monitored by a carotid arterial line using a physiologic pressure transducer (Berg GmbH, Dirchseeon/Eglharting, Germany). Ventilatory parameters were adjusted on the basis of blood gas values (pO2, 148.5 ± 16.4; pCO2, 41.1 ± 3.4; and pH, 7.43 ± 0.08) (AVL 995-Hb, AVL List GmbH, Graz, Austria) and oxygen saturation using a hemoximeter (OSM-2, Radiometer, Copenhagen, Denmark). The data relating to arterial blood gas and oxygen saturation during the entire period of investigation corresponded to the respective physiologic values. No significant changes of pH values were detected in SHAM1, SHAM2.5, and IR1 groups through all experiments. In IR2.5 group, however, pH decreased from the baseline of 7.43 ± 0.08 to 7.25 ± 0.10 at the end of reperfusion (P < .005). This can be explained as a result of anaerobic glycolysis and production of lactate with subsequent development of acidosis during normothermic ischemia. Ringer's solution (Fresenius Pharma, Graz/Linz Austria) was infused at a dose of 0.2 mL/kg/min through the auricular vein. Body temperature was monitored online by a rectal probe and maintained at 37.5 ± 0.3°C by an infrared light and heating mat. For blood sampling, a catheter was placed by the internal iliac vein into the distal inferior caval vein of the animals. Venous blood samples (3 mL each) were obtained before ischemia (BI), at the end of ischemia (EI), and 2 hours after reperfusion (R2). To compensate for the blood loss, an adequate quantity of isotonic sodium chloride (3 mL) was infused after each sampling. Bilateral hind-limb ischemia was induced by established techniques.7 Common femoral arteries were clamped in the groins with microclips, and collateral blood flow was occluded by wrapping a rubber arterial tourniquet (Stripp-Quick, KaWe, Stuttgart, Germany) around each thigh at the proximal third of the leg. Occlusion of blood flow was confirmed using a blood perfusion monitor (Laserflo BPM2, Vasamedics, St Paul, Minn). Both limbs were rendered ischemic for either 1 (IR1 group) or 2.5 (IR2.5 group) hours. For reperfusion, the clamps and tourniquets were released. Reperfusion was verified by parameters on the basis of restoration of pulsatile blood flow in the femoral arteries and on the data of the blood perfusion monitor. The sham operation consisted of groin incisions and dissection of femoral blood vessels. After completion of the experiments, the animals were killed by intravenous administration of an overdose of potassium chloride. Morphometric analysis At the 3 time points of investigation (BI, EI, R2), biopsies were obtained from the great adductor muscles of the right hind limbs and were immediately immersed for quick freezing at −70°C in a 2-methylbutane solution (Uvasol, Merck, Darmstadt, Germany) for 2 minutes. Samples were stored at −80°C until sectioned. Transverse cryosections 10 μm-thick (Kryostat 1720, Leitz, Germany) were stained for actomyosin adenosine thriphosphatase activity at pH 4.3 and 10.2. After the different stainings were compared by an unbiased observer, the sections stained at pH 4.3 were examined by light microscopy (Axiomat, Zeiss, Oberkochen, Germany). The morphometric measurements and fiber and microvessel counts were performed with a pen connected to a personal computer using a semiautomatic image analyzer (Lucia_M, Nikon Laboratory Imaging, Prague, Czech Republic) at 100× (fibers) or 1000× (microvessels) magnification. Fibers were counted per 3 random fields in each section and the fraction of muscle interfiber area (percentage of cross-section area composed of interfiber space) determined. Fifty microvessels were measured per random field in each section, and the microvessel cross-section area (in μm2) was determined. Electron microscopy Muscle samples were immersion-fixed slightly stretched in 3% glutardialdehyde and then postfixed in 1% osmium tetroxide, both with 0.1 mol/L cacodylate buffer (pH 7.4). Samples were dehydrated in ethanol and embedded in epoxy resin (Epon). Before preparing the ultrathin sections, semithin sections (2 μm) of transverse and longitudinal muscles were prepared and stained for toluidine blue. Ultrathin cross-sections and longitudinal sections (90 nm) of the Epon-embedded muscle samples were mounted on 200-mesh copper grids, stained in a 2% uranyl acetate solution followed by 0.4% lead citrate in 0.1 mol/L sodium hydroxide, and examined under a transmission electron microscope (EM 10, Zeiss, Germany). Assessment of plasma lipid peroxide Venous blood was collected in EDTA tubes at the time intervals mentioned above and immediately centrifuged (3000× g for 7 minutes at +20°C). Subsequently, the plasma was separated and stored at −80°C until assayed. Hemolytic plasma samples (1 sample from IR1 and 1 from IR2.5 groups) were excluded. Lipid peroxidation was assessed by measuring malondialdehyde levels (MDA) in plasma (pMDA) as described by Wong et al.16 The value obtained by this method may reflect some combination of free and bound MDA originally present in the sample plus MDA formed from oxidized lipids during the procedure. This method, therefore, provides a measure of lipid peroxidation within the sample in terms of MDA equivalents. Assessment of pTau Heparinized venous blood samples were centrifuged at 3000 g for 7 minutes. The plasma was separated and deproteinized with sulfosalicylic acid (30%) containing 1 mmol/L β-(2-thienyl) ± alanine as an internal standard. After centrifugation, an aliquot of the supernatant was diluted in membrane-filtered water, derivatized with o-phthaldialdehyde, and immediately analyzed for pTau. Assessment of taurine levels in skeletal muscle tissue Total taurine Muscle biopsies obtained from the great adductor muscle of the right hind limb at time points BI and R2 were immediately wet-weighted and homogenized in a glass container filled with 500 μL of ice-cold sufosalicylic acid (4%) containing β-(2-thienyl) ± alanine as an internal standard. The homogenate was cooled on ice for 30 minutes, then centrifuged at 12,000× g for 5 minutes. An aliquot of the supernatant was derivatized with o-phthaldialdehyde and immediately analyzed. Total taurine concentrations in skeletal muscles were expressed as μmol/L of analyzed liquid. The taurine contents in skeletal muscles were derived from the total taurine concentration and the fat-free dry mass of skeletal muscle sample (after extraction by petrol ether). The muscular taurine contents were expressed as mmol/100 g of fat-free dry mass. iTau This parameter was obtained by subtracting the extracellular from the total muscular taurine concentration. To this end, the extracellular taurine concentration in muscle tissue was assumed to be equal to the plasma level, and the intracellular water volume needed to calculate the parameter was quantitated by the chloride method.17 The iTau level in skeletal muscle tissue was expressed as mmol/L of intracellular fluid. Taurine was detected by fluorescence (Excitation wavelength: 330 nm; Emission wavelength: 440 nm) and separated from other amino acid compounds using high-performance liquid chromatography on a 3 μm, 125 × 4.6 mm ODS Hypersil column (Bischoff, Leonberg, Germany). Assessment of plasma CK and LDH levels The plasma levels of CK and LDH were measured in blood plasma samples using laboratory analyser (904, Hitachi, Japan). Functional assessment of limb skeletal muscles 3 weeks after operation In 5 animals of each IR1 and IR2.5 groups and in 5 control untreated animals the functional assessment of the rectus femoris muscle was performed 3 weeks after operation. The animals had free access to food and water. Postoperative analgesia was performed for 3 days using buprenorphin hydrochlorid 3 μg/kg body weight (Temgesic, Boehringer Mannheim, Vienna). After 3 weeks, the animals were anesthetized with intravenous ketamine and xylazine and intubated. Anesthesia was maintained with a mixture of halothane, nitrous oxide, and oxygen. Maximal tetanic tension Maximal tetanic tension (MTT) of the rectus femoris muscle was investigated according to Koller et al.18 In both groins the femoral nerves were exposed. In both hind limbs the rectus femoris muscles were exposed and free dissected. Only the neuromuscular pedicle and the origin of the muscle were preserved. The patellar ligament was transected and the patella was connected to a force transducer (Hottinger Baldwin Messtechnik, Darmstadt, Germany). The pelvic bone and the femur were rigidly immobilized in a frame. For the electrostimulation, bipolar electrode was connected to the distal part of the femoral nerve on the left side and electrostimulation was performed. After the test was completed, the same procedure was applied on the contralateral side. Isometric tetanic contractions of both rectus femoris muscles were registred at supramaximal stimulation under a constant resting tension of 4 N. The following stimulation parameters were used: bipolar burst stimulation with duration 25 ms; pulse width 1001 μs; frequency 67 Hz; number of impulses pro burst, 40. The highest achievable tension was used for statistical analysis. Statistical analysis Results are given as mean values and SD. The studied groups and time points were compared with each other using blocked analysis of variance. Means were considered statistically different at P < .05. All statistical analysis was exploratory rather than confirmatory and was performed using SAS statistical software (SAS 1990, SAS/STAT user's guide, v. 6, SAS Institute, Cary, NC). Results Histology The histologic sections from both sham groups had the typical appearance of normal muscle tissue featuring distinct fascicles of polygonal muscle fibers with subsarcolemmal myonuclei and a normal interstitium (Fig 1, A, and 2, A).
The IR1 and IR2.5 groups did not show any discernible morphologic changes at EI. The interfiber and microvessel cross-section areas did not significantly change as compared with the corresponding sham groups (Tables I and II).
| | |  | Groups | SHAM1 (n = 6) | IR1 (n = 7) | SHAM2.5 (n =7) | IR2.5 (n = 8) | |
|---|
| BI | 13 ± 2 | 14 ± 1 | 13 ± 2 | 13 ± 2 | |
| EI | 15 ± 1 | 14 ± 1 | 14 ± 1 | 15 ± 1 | |
| R2 | 14 ± 1 | 17 ± 1* | 14 ± 1 | 28 ± 4** | |
| | | | | |
| | | | Groups | SHAM1 (n = 6) | IR1 (n = 7) | SHAM2.5 (n = 7) | IR2.5 (n = 8) | |
|---|
| BI | 20 ± 1 | 19 ± 1 | 19 ± 1 | 19 ± 1 | |
| EI | 20 ± 1 | 19 ± 1 | 18 ± 2 | 19 ± 1 | |
| R2 | 20 ± 1 | 19 ± 1 | 19 ± 1 | 13 ± 1* | |
| | | | | |
After reperfusion, the IR1 group did show mild interstitial edema (Fig 1, B) as indicated by a 21% increase in interfiber area (Table I), whereas the microvessel cross-section area essentially remained unchanged (Table II). Muscle fibers showed no discernible histologic changes (Fig 2, B). Severe morphologic changes after reperfusion did occur in the IR2.5 group, including membrane ruptures with dissolution of myofibrils (Fig 2, C). Likewise, the IR2.5 group showed marked interstitial edema (Fig 1, C), as reflected in a 112% increase of the interfiber area. Interstitial edema formation after reperfusion was significantly higher than in the IR1 group (P < .005) (Table I). Moreover, the reperfused muscles in this group were affected by microvascular constriction, as evidenced by a 33% reduction of the microvessel cross-section area (P < .0005) (Table II). Electron microscopy Electron micrographs of the ultrathin cross-sections obtained BI, at EI, and after reperfusion demonstrated that the morphology of skeletal muscles in both sham groups remained normal throughout the observation period (Fig 3, A).
The IR1 group was morphologically normal at EI. After reperfusion, the normal but also swollen mitochondria appeared in muscle fibers although no sarcolemmal damage was noticed (Fig 3, B). The IR2.5 group displayed some mildly swollen mitochondria at EI, followed by pronounced structural lesions in most muscle fibers after reperfusion (Fig 3, C and D). In particular, the sarcolemma were severely damaged and the basal lamina interrupted. Myofibrils were frequently disoriented and partially dissolved. Mitochondria were swollen, damaged, and accumulated at the periphery of the muscle fibers. The cristae in these mitochondria were degenerated and the membranes characterized by discontinuities. The sarcoplasmic reticulum was extensively dilated; nuclei were pycnotic and showed peripheral accumulation of chromatin. pMDA concentrations Neither the sham groups nor the IR1 group showed significant changes in basal pMDA (0.67 ± 0.11 μmol/L). In the IR2.5 group, by contrast, pMDA was significantly higher than in the corresponding SHAM2.5 group at the end of reperfusion (0.81 ± 0.14 vs 0.57 ± 0.11 μmol/L, P < .05). Tissue taurine and iTau concentrations Basal concentrations of tissue taurine (250 ± 90 μmol/100 g fat-free dry mass) and iTau (1070 ± 400 μmol/L intracellular fluid) never revealed significant changes in any of the study groups. Plasma concentrations of CK and LDH No significant changes of plasma concentrations of CK and LDH were observed in the sham groups or the IR1 group. In the IR2.5 group, CK and LDH levels were significantly higher than in the corresponding SHAM2.5 group R2 (P < .005 and P < .05, respectively) (Table III). Increase of plasma CK concentrations from the baseline was about 4-fold, whereas the increase of LDH was in the same magnitude as for pTau. Functional assessment of limb skeletal muscle 3 weeks after operation MTT MTT of the rectus femoris muscle in the control untreated animals was 52.52 ± 3.59 N (n = 5). MTT of the same muscle in the IR1 group (45.04 ± 8.67 N, n = 5) did not differ statistically from the corresponding control value. However, significant reduction of MTT of the rectus femoris muscle was observed in the animals of IR2.5 group (27.36 ± 6.74 N, n = 5) as compared with the control value (P < .005). Discussion Taurine is a nonessential amino acid that is present at high concentrations in excitable and membrane-rich tissues.17 In mammals it is supplied by 2 pathways: endogenous biosynthesis of cysteine-sulfinic acid19 and exogenous dietary supplementation20 or parenteral nutrition.21 The exact physiologic role of taurine remains to be established, although various functions have been described. Taurine has been shown to be instrumentally involved in the electrophysiologic functions of muscle cell membranes,22, 23 to regulate calcium ion (Ca2+) homeostasis and cell volume by osmotic properties,24, 25 and to stabilize muscle cell membranes.24 Skeletal muscle tissue constitutes a large pool of iTau,10 which is maintained by active uptake, endogenous synthesis, or both and is located in myofilaments, sarcoplasmic membranes, reticula, matrix, and, to a lesser extent, in mitochondria and capillaries.26 The entry of taurine into the cells is by means of a specific sodium dependent Na+/Cl− coupled transport.11 Taurine concentration gradients across cell membranes are extremely high: gradients of 500:1 are maintained in retinal or brain cells,27 and a value as high as 300:1 has been reported for human muscle tissue.12 These high gradients are maintained by the fact that the taurine is rendered water-soluble and high lipophobic by the action of its β-amino acid. This results in an extremely low diffusion of taurine through lipophilic membranes. Therefore, if membranes are disrupted,28 or sodium dependent transporter is compromised29 iTau may leak into the bloodstream and give rise to increased pTau. In our model of I/R injury to skeletal muscles, increased pTau were only observed with the longer ischemic interval (2.5 hours) followed by 2 hours of reperfusion, whereas the shorter ischemic interval (1 hour) did not raise pTau. These differences in pTau reflected morphologic and biochemic changes in reperfused skeletal muscles after brief versus prolonged ischemia. It is well known that I/R results in a depletion of cellular energy stores of high energy phosphate bonds: adenosine triphosphate and phosphocreatine, and glycogen stores. It was shown that energy metabolism in skeletal muscle was influenced by 2.5 hours of ischemia, decreasing intracellular phosphocreatine by 65%, whereas adenosine triphosphate remained stable. After reperfusion, a further depletion of phosphocreatine was noticed that was accompanied by a depletion of adenosine triphosphate by 50% from the initial level.30 The consequences of such depletion are a compromised active cellular transport system and a loss of normal cell gradients. The mild interstitial edema formation observable in our IR1 group did not involve any irreversible structural changes after 1 hour of ischemia and R2. In particular, the cell membranes were intact. This is in accordance with the findings of Rácz et al,31 who described cloudy swelling and edematous perimysium of skeletal muscles after 1 hour of ischemia and R2. Likewise, the IR1 group did not reveal any abnormalities in intermuscular microcirculation and involved no changes in the size of microvessels. In contrast, reperfusion after 2.5 hours of ischemia in the IR2.5 group led to severe interstitial edema formation and significant damage to muscle fibers. Microvessel size was significantly reduced in the IR2.5 group and was associated with a no-reflow phenomenon, ie, a persistent decrease in blood flow after the initial hyperfusion phase.2 In addition, the IR2.5 group was characterized by mitochondrial swelling, loss of mitochondrial cristae, and dilatation of sarcoplasmic reticula. Similar results were described in a previous design involving 2 to 3 hours of ischemia and 30 minutes of reflow.32 Sternbergh et al33 reported ultrastructural damage to skeletal muscles after 2 hours, but not after 1 hour, of ischemia and 1 hour of reperfusion. Our morphologic evidence of muscle tissue damage (particularly to the cell membranes) after reperfusion in the IR2.5 group coincided with increased pMDA, demonstrating an enhanced process of membrane lipid peroxidation by excessive formation of oxygen free radicals.34 Considerable rises in MDA were described previously using 2 hours of ischemia and 1 hour of reperfusion.35 In our experiment, 1 hour of ischemia and 2 hours of reperfusion did not give rise to increased pMDA, reflecting the finding of intact cell membranes at the ultrastructural level. Our finding of significant interstitial edema formation and membrane rupture resulting from excessive lipid peroxidation coincided with a significant rise in pTau in the IR2.5 group, ie, after 2.5 hours of ischemia and R2. Lipid peroxidation and membrane rupture affected cell transport functions and, by implication, the permeability to taurine. The breakdown of muscle cell membranes caused large amounts of iTau to be released to the interstitium that raised pTau whereas iTau and tissue taurine concentrations remained unchanged. The fact that we observed no significant depletion of iTau may be a result of its extremely high concentrations, but it is also known that changed plasma concentrations of amino acids are not automatically reflected in muscle tissue.12 The time pattern of the blood taurine concentrations was qualitatively similar to that of the CK and LDH activities, ie, all 3 parameters demonstrated increased values after reperfusion of skeletal muscle after 2.5 hours but not 1 hour of ischemia influence. However, increase of the CK activities was greater (4.2 times) than the rise in taurine (2.6 times) and LDH (2.5 times) concentrations. Pronounced cell damage in IR2.5 group after reperfusion resulted in impaired muscle function 3 weeks after I/R injury: muscle MTT was reduced 2 times as compared with control value. Contrarily, no significant muscle function impairment was noticed in group IR1. Consequently, taurine leakage from cells and subsequent increase of pTau after ischemia may be considered as prognostically not favorable in terms of reversibility of organ function. Our study demonstrates that pTau correlates with different ischemic intervals involved in I/R injury of skeletal muscle. Increased values were observed after 2.5 hours, but not after 1 hour of ischemia followed by 2 hours of reperfusion, which reflected the presence or absence of severe damage of muscle cell membranes and late muscle dysfunction. We therefore conclude that pTau may be a sensitive marker of I/R injury to skeletal muscle, providing useful diagnostic and prognostic information. Acknowledgements We are indebted to Rupert Koller, MD, for assistance in performing muscle functional measurements. References 1.
1
Nanobashvili J, Neumayer C, Fuegl A, Sporn E, Prager M, Polterauer P, et al.
Ischemia/reperfusion injury of skeletal muscle: mechanisms, morphology, treatment strategies, and clinical applications.
European Surgery. 2002;34:83–89. 2.
2
Huk I, Brovkovych V, Nanobashvili J, Weigel G, Neumayer C, Partyka L, et al.
Bioflavonoid quercetin scavenges superoxide and increases nitric oxide concentration in ischaemia-reperfusion injury: an experimental study.
Br J Surg. 1998;85:1080–1085. MEDLINE |
CrossRef
3.
3
Freischlag JA, Hanna D.
Superoxide anion release (O2−) after ischaemia and reperfusion.
J Surg Res. 1991;50:565–568. MEDLINE |
CrossRef
4.
4
Huk I, Brovkovych V, Nanobashvili J, Neumayer C, Polterauer P, Prager M, et al.
Prostaglandin E1 reduces ischemia/reperfusion injury by normalizing nitric oxide and superoxide release.
Shock. 2000;14:234–242. MEDLINE |
CrossRef
5.
5
Idström JP, Soussi B, Elander A, Bylund-Fellenius AC.
Purine metabolism after in vivo ischemia and reperfusion in rat skeletal muscle.
Am J Physiol Heart Circ Physiol. 1990;258:H1668–H1673. 6.
6
Roth E, Manhart N, Punz A.
Antioxidative abwehrmechanismen während systemischer inflammation.
Intensivmed. 1998;35:95–105. 7.
7
Huk I, Nanobashvili J, Neumayer C, Punz A, Mueller M, Afkhampour K, et al.
L-Arginine treatment alters the kinetics of nitric oxide and superoxide release and reduces ischemia/reperfusion injury in skeletal muscle.
Circulation. 1997;96:667–675. MEDLINE 8.
8
McCord JM.
Oxygen-derived free radicals in postischemic tissue injury.
N Engl J Med. 1985;312:159–163. MEDLINE 9.
9
Hernández-Maldonado JJ, Teehan E, Franco CD, Durán WN, Hobson RW.
Superoxide anion production by leukocytes exposed to post-ischemic skeletal muscle.
J Cardiovasc Surg (Torino). 1992;33:695–699. MEDLINE 10.
10
Turner O, Phoenix J, Wray S.
Developmental and gestational changes of phospho-ethanolamine and taurine in rat brain, striated and smooth muscle.
Exp Physiol. 1994;79:681–689. MEDLINE 11.
11
Huxtable RJ.
Physiological actions of taurine.
Physiol Rev. 1992;72:101–163. MEDLINE 12.
12
Roth E, Zöch G, Schulz F, Karner J, Mühlbacher F, Hamilton G, et al.
Amino acid concentrations in plasma and skeletal muscle of patients with acute hemorrhagic necrotizing pancreatitis.
Clin Chem. 1985;31:1305–1309. MEDLINE 13.
13
Suleiman MS, Fernando HC, Dihmis WC, Hutter JA, Chapman RA.
A loss of taurine and other amino acids from ventricles of patients undergoing bypass surgery.
Br Heart J. 1993;69:241–245. MEDLINE |
CrossRef
14.
14
Lombardini JB, Cooper MW.
Elevated blood taurine levels in acute and evolving myocardial infarction.
J Lab Clin Med. 1981;98:849–859. MEDLINE 15.
15
Kennergren C, Mantovani V, Lonnroth P, Nystrom B, Berglin E, Hamberger A.
Extracellular amino acids as markers of myocardial ischemia during cardioplegic heart arrest.
Cardiology. 1999;91:31–40. 16.
16
Wong SHY, Knight JA, Hopfer SM, Zaharia O, Leach CN, Sunderman FW.
Lipoperoxides in plasma as measured by liquid-chromatographic separation of malondialdehyde-thiobarbituric acid adduct.
Clin Chem. 1987;33:214–220. MEDLINE 17.
17
Wright CE, Tallan HH, Lin YY, Gaull GE.
Taurine: biological update.
Annu Rev Biochem. 1986;55:427–453. MEDLINE 18.
18
Koller R, Rab M, Todoroff BP, Neumayer Ch, Haslik W, Stöhr HG, et al.
The influence of the graft length on the functional and morphological results after nerve grafting: an experimental study in rabbits.
Br J Plast Surg. 1997;50:609–614. Abstract |
Full-Text PDF (1139 KB)
|
CrossRef
19.
19
Huxtable RJ.
Taurine in the central nervous system and the mammalian actions of taurine.
Prog Neurobiol. 1989;32:471–533. 20.
20
Gaull GE.
Taurine as a conditionally essential nutrient in man.
J Am Coll Nutr. 1986;5:121–125. MEDLINE 21.
21
Geggel HS, Ament ME, Heckenlively JR, Martin DA, Kopple JD.
Nutritional requirement for taurine in patients receiving long-term parenteral nutrition.
N Engl J Med. 1985;312:142–146. MEDLINE 22.
22
Airaksinen EM, Paljärvi L, Partanen J, Collan Y, Laakso R, Pentikäinen T.
Taurine in normal and diseased human skeletal muscle.
Acta Neurol Scand. 1990;81:1–7. MEDLINE 23.
23
Pierno S, De Luca A, Camerino C, Huxtable RJ, Camerino DC.
Chronic administration of taurine to aged rats improves the electrical and contractile properties of skeletal muscle fibers.
J Pharmacol Exp Ther. 1998;286:1183–1190. MEDLINE 24.
24
Huxtable R, Bressler R.
Effect of taurine on a muscle intracellular membrane.
Biochim Biophys Acta. 1973;323:573–583. MEDLINE 25.
25
Schaffer SW, Azuma J, Madura JD.
Mechanisms underlying taurine-mediated alterations in membrane function.
Amino Acids. 1995;8:231–246.
CrossRef
26.
26
Terauchi A, Nagata T.
Observation on incorporation of 3H-taurine in mouse skeletal muscle cells by light and electron microscopic radioautography.
Cell Mol Biol (Noisy-le-grand). 1993;39:397–404. MEDLINE 27.
27
Pasantes-Morales H, Cruz C.
Taurine: a physiological stabilizer of photoreceptor membranes.
Prog Clin Biol Res. 1985;179:371–381. MEDLINE 28.
28
Banks MA, Porter DW, Martin WG, Castranova V.
Effects of in vitro ozone exposure on peroxidative damage, membrane leakage, and taurine content of rat alveolar macrophages.
Toxicol Appl Pharmacol. 1990;105:55–65.
CrossRef
29.
29
Saransaari P, Oja SS.
Mechanisms of ischemia-induced taurine release in mouse hippocampal slices.
Brain Res. 1998;807:118–124. MEDLINE |
CrossRef
30.
30
Neumayer C, Nanobashvili J, Fügl A, Blumer R, Punz A, Prager M, et al.
α-Tocopherolpretreatment reduces ischemia/reperfusion injury in rabbit skeletal muscle.
Eur Surg. 2002;34:90–94. 31.
31
Rácz IB, Illyés G, Sarkadi L, Hamar J.
The functional and morphological damage of ischemic reperfused skeletal muscle.
Eur Surg Res. 1997;29:254–263.
CrossRef
32.
32
Santavirta S, Luoma A, Arstila AU.
Ultrastructural changes in striated muscle after experimental tourniquet ischaemia and short reflow.
Eur Surg Res. 1978;10:415–424.
CrossRef
33.
33
Sternbergh WC, Adelman B.
The temporal relationship between endothelial cell dysfunction and skeletal muscle damage after ischemia and reperfusion.
J Vasc Surg. 1992;16:30–39. Abstract | Full Text |
Full-Text PDF (1755 KB)
34.
34
Kawasaki S, Sugiyama S, Ishiguro N, Ozawa T, Miura T.
Implication of superoxide radicals on ischemia-reperfusion induced skeletal muscle injury in rats.
Eur Surg Res. 1993;25:129–136.
CrossRef
35.
35
Lee YH, Wei FC, Lee J, Su MS, Chang YC.
Effect of postischemic reperfusion on microcirculation and lipid metabolism of skeletal muscle.
Microsurgery. 1995;16:522–527. MEDLINE |
CrossRef
Departments of Vascular Surgery, Anatomy, Surgical Research Laboratories, Ludwig Boltzmann Research Institute for Interdisciplinary Vascular Medicine, Department of Medical Computer Sciences, University of Vienna Medical School, Vienna, Austria; and Department of Chemistry and Biochemistry, Ohio University, Athens, Ohio ☆ Supported in part by a grant from Public Health Service (HL 55397). ☆☆ Reprint requests: Joseph Nanobashvili, MD, Department of Vascular Surgery, University of Vienna Medical School, Waehringer Guertel 18-20, A-1090, Vienna, Austria. ★ 0039-6060/2003/$30.00 + 0 PII: S0039-6060(02)21671-2 doi:10.1067/msy.2003.65 © 2003 Published by Elsevier Inc. | |
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