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1 Department of Medical Anatomy, The Panum Institute, University of Copenhagen, Blegdamsvej 3, DK-2200, Copenhagen, Denmark 2 Department of Infectious Diseases and the Copenhagen Muscle Research Centre, Rigshospitalet, Blegdamsvej 9, DK-2100, Copenhagen, Denmark 3 Institute of Neurosciences and Department of Cellular Biology, Physiology and Immunology, Animal Physiology Unit, Faculty of Sciences, Autonomous University of Barcelona, Barcelona, Spain 08193
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Abstract |
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(Received 18 October 2004;
accepted after revision 20 December 2004; first published online 7 January 2005)
Corresponding author M. Penkowa: Department of Medical Anatomy, The Panum Institute, University of Copenhagen, Blegdamsvej 3, DK-2200, Copenhagen, Denmark. Email: m.penkowa{at}mai.ku.dk
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Introduction |
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MT-I + II are significant antioxidant factors, which also have key anti-apoptotic functions, whereby MT-I + II protect tissues and cells during various pathological conditions (Aschner et al. 1997; Viarengo et al. 2000; Penkowa et al. 2000, 2001; Hidalgo et al. 2001, 2002; Giralt et al. 2002b; Penkowa, 2003). MT-I + II act by inhibiting reactive oxygen species (ROS)-induced degradation of DNA and tissue damage (Abel & de Ruiter, 1989; Cai et al. 2000; Kling & Olsson, 2000; Rana & Kumar, 2000), and by scavenging and inhibiting the formation of ROS (Thornalley & Vasak, 1985; Schwarz et al. 1994, 1995; Lazo et al. 1998; Viarengo et al. 2000). MT-I + II have important roles during oxidative stress by interaction with reduced glutathione (GSH)(Jiang et al. 1998), and by inhibiting GSH depletion (Haidara et al. 1999). Moreover, MT-I + II can antagonize the deleterious effects of oxidative stress on catalase (Haidara et al. 1999).
It has been shown that genetic MT-I overexpression as well as infusion of MT-II protects against oxidative stress and tissue injury (Schwarz et al. 1994, 1995; Pitt et al. 1997; Suzuki et al. 2000; Giralt et al. 2002b; Penkowa et al. 2002, 2003a). In addition, genetic MT-I + II deficiency leads to a dramatic increase in oxidative stress and cellular damage (Lazo & Pitt, 1995; Zheng et al. 1996; Carrasco et al. 2000; Suzuki et al. 2000; Penkowa et al. 2000, 2001; Giralt et al. 2002a).
A substantial amount of data indicates that exercise is associated with an increase in the production of free radicals and ROS by skeletal muscle (Davies et al. 1982; Jackson et al. 1985). This increase appears to occur because a proportion of the molecular oxygen used in normal respiration undergoes a one-electron reduction to generate superoxide radicals (Boveris et al. 1972). It has been estimated that exercise can increase oxygen utilization 200-fold above resting levels in active muscle fibres and superoxide production appears to increase with this large increase in oxygen flux through muscle mitochondria during exercise. The potential consequences of the exercise-induced increase in oxidative stress have been addressed in a number of studies. Data have been presented suggesting that oxidative stress plays a role in the muscle fatigue or damage (Andrade et al. 1998) that accompanies some forms of exercise, although these roles remain contentious (Jackson, 1996; Patwell et al. 2004). Some studies report that an acute bout of exercise increases the activities of superoxide dismutase (SOD), glutathione peroxidase, glutathione reductase and catalase in skeletal muscle of rats (Ji, 1993). In humans, exercise training has been reported to increase skeletal muscle SOD activity (Jenkins et al. 1984) and the activities of various protective enzymes in blood (Robertson et al. 1991). Oxidative and other stresses to cells are also known to induce the formation of heat shock proteins (HSPs). HSPs represent important components of the cellular protective response and data also indicate that an increase in muscle HSP content occurs after exercise in rodents (Salo et al. 1991; Kelly et al. 1996; Hernando & Manso, 1997; McArdle et al. 2004) and in humans (Febbraio & Koukoulas, 2000; Moseley, 2000; Khassaf et al. 2001; Febbraio et al. 2002). These proteins are thought to facilitate repair from injury and to aid adaptation and remodelling of the cell to prevent the damage from recurring after a repeat of the same stress (Patwell et al. 2004; McArdle et al. 2004). Thus, intramyocelluar expression of some HSPs may counteract the muscle-damaging effects of oxidative stress.
Given that MT-I + II are induced by oxidative stress and protect against tissue damage, we suggest that these proteins might represent an alternative pathway whereby muscle fibres are protected from oxidative stress-induced damage. Little information exists with regard to MT and physical activity. In 1978 it was shown that MT synthesis increased in the rat liver after strenuous exercise (Oh et al. 1978), whereas a later study found that chronic exercise led to a decrease in the amounts of MT-I + II in aortic vessel tissue (Bobillier et al. 2001). Here we demonstrate that physical exercise, and the associated muscular oxidative stress, is accompanied by increased MT-I + II expression within human skeletal muscle fibres.
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Methods |
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One week before the first experimental day, subjects performed a maximal oxygen uptake test (VO2,max) on an ergometer bicycle. Measurement of VO2,max was performed on an electrically braked cadence-independent cycle ergometer (Monark 839E, Monark Ltd, Varberg, Sweden). Subjects cycled for 3 min at 100 W as a warm-up followed by cycling at progressively higher work rates, increasing 50 W every 3 min for 9 min and then increasing the workload by 25 W every minute until subjects were unable to maintain a cadence of 60 r.p.m. Expired oxygen (VO2) and carbon dioxide (VCO2) concentrations were recorded online. Median VO2,max was 46.7 ml kg–1 min–1 (range, 41–68). On the experimental day, subjects arrived at 08.00 h after an overnight fast and were randomly assigned to either exercise or rest. There was no difference between the two groups regarding age, weight, height or VO2,max. Subjects performed 3 h of cycling (n = 12) at 60% VO2,max followed by 6 h of recovery. The following day they reported to the laboratory after an overnight fast. Prior to the exercise (0), immediately after the exercise (3 h), and at 4.5, 6, 9 and 24 h, muscle biopsies were obtained from the vastus lateralis using the percutaneous needle biopsy technique with suction. Control subjects rested in the laboratory for 9 h (n = 6), reported to t he laboratory the day after and had biopsies collected at the same time points.
Tissue processing
Muscle tissue was cut in 6-µm consecutive sections on a cryostat, and the sections were immediately collected on glass slides, to be used for general histology and immunohistochemistry.
Sections were preincubated in 0.5% H2O2 in Tris-buffered saline (TBS)/Nonidet (0.05 M Tris, pH 7.4 and 0.15 M NaCl with 0.01% Nonidet P-40; Sigma-Aldrich, St Louis, MO, USA) for 15 min at room temperature (20°C) to quench endogenous peroxidase. Afterwards, sections were washed in TBS/Nonidet three times for 5 min. Sections were also preincubated with 10% normal goat serum (In Vitro, DK) in TBS/Nonidet for 30 min at room temperature in order to block non-specific binding.
Histochemistry
Haematoxylin and eosin (HE) staining of the sections was performed according to standard procedures. Also, myofibrillar ATPase staining with preincubation at pH 4.6 was used to identify muscle fibre types (Brooke & Kaiser, 1970).
Immunohistochemistry
The sections stained immunohistochemically were always processed simultaneously and under the same laboratory conditions.
Sections were incubated overnight at 4°C with one of the following primary antibodies: mouse anti-horse MT-I + II diluted 1: 50 (Dakopatts, DK); rabbit anti-rat MT-I + II diluted 1: 500 (Gasull et al. 1993); rabbit anti-malondialdehyde (MDA) diluted 1: 100 (Alpha Diagnostics, San Antonio, TX, USA) (as a marker of lipid peroxidation/oxidative stress); and rabbit anti-nitrotyrosine (NITT) diluted 1: 200 (Alpha Diagnostics) (as a marker of peroxynitrite-induced protein nitration/oxidative stress). These primary antibodies were detected using biotinylated goat anti-mouse IgG diluted 1: 200 (Sigma-Aldrich) or biotinylated mouse anti-rabbit IgG diluted 1: 400 (Sigma-Aldrich) for 30 min at room temperature followed by streptavidin-biotin-peroxidase complex (StreptABComplex/HRP, Dakopatts, DK) prepared according to the manufacturer's recommended dilutions for 30 min at room temperature. Afterwards, sections were incubated with biotinylated tyramide and streptavidin-peroxidase complex (NEN, Life Science Products, Boston, MA, USA) pre-pared according to the manufacturer's recommendations. The immunoreaction was visualized using 0.015% H2O2 in 3,3-diaminobenzidine-tetrahydrochloride (DAB)/TBS for 10 min at room temperature.
In order to evaluate the extent of non-specific binding in the immunohistochemical experiments, control sections were incubated in the absence of primary antibody or in the absence of secondary antibody or in the blocking serum. Results were considered only if these controls were negative.
In order to exclude staining due to endogenous biotin, we have pretreated sections sequentially with 0.01–0.1% avidin (Sigma-Aldrich) followed by 0.001–0.01% biotin (Sigma-Aldrich), each step for 20 min at room temperature, before the immunohistochemical analysis was performed. Comparing our immunohistochemical stainings with/without specific biotin blocking showed that in the tissue, muscular endogenous biotin is unlikely to induce false-positive immunostaining by binding to the streptavidin. In order to control the specificity of the primary antibodies we have pre-absorbed the primary antibodies with their corresponding antigenic proteins. For this purpose we have used MT-I and MT-II (both from Sigma-Aldrich) proteins for the anti-MT-I + II antibodies, nitrosylated/nitrated proteins (Alpha Diagnostics) for the anti-NITT antibodies and MDA-ovalbumin (Alpha Diagnostics) for the anti-MDA antibodies. Results were considered only if this pre-absorbtion of primary antibodies resulted in negative immunostaining.
For the simultaneous examination and recording of the staining, a Zeiss Axioplan2 light microscope was used.
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PCR |
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Reverse transcription. Total RNA (1 µg) was reverse transcribed in a 50-µl reaction according to the manufacturer's instructions (Applied Biosystems, Taqman reverse transcription reagents) with the use of Oligo dT primers. The reactions were run in a Perkin Elmer GeneAmp PCR system 9700 at 25°C for 10 min, 48°C for 30 min, and 95°C for 5 min.
Analysis of gene expression levels in muscle tissue. MT-II primers and Taqman probe were designed using Primer Express 2.0. The primers and probe spanned an intron boundary to prevent amplification of genomic DNA. To ensure the specificity of the primers and probe a gel was run, where a single band of the correct size of 103 bp was obtained. MT-II: forward primer, 5'-CGCCATGGATCCCAACT-3'; reverse primer, 5'-GCAGCTTTTCTTGCACGAAGT-3'; Taqman probe, 5'-CCGCCGGTGACTCCTGCACCT-3'. The reaction conditions for the MT-II real-time PCR were: forward primer, 50 nM, reverse primer 900 nM and Taqman probe 200 nM.
Samples were analysed for MT-II mRNA levels by real-time PCR using an ABI PRISM 7900 sequence detector (PE Biosystems). Samples were run in triplicate under standard real-time PCR conditions: 50°C for 2 min, 95°C for 10 min followed by 50 cycles of 95°C for 15 s and 60°C for 1 min. The gene expression levels were normalized to the housekeeping gene glyceraldehyde-3 phosphate dehydrogenase (GAPDH; obtained from Applied Biosystems). All reactions were run under singleplex conditions. Data were quantified and normalized using the standard curve method. A linear correlation of 0.99 was obtained for the amplification of both MT-II and GAPDH cDNA.
Statistics
Data for MT-II/GAPDH mRNA ratios were normally distributed after log-transformation. A two-way RM-ANOVA was used to detect changes over time or between groups. Student–Newman–Keul's t test for post hoc analysis was used to detect changes over time from resting values or differences between groups. P < 0.05 was considered significant. Statistical calculations were performed using Sigma Stat 3.0 (SPSS Inc., Chicago, IL, USA). Data are presented as geometric mean ± S.E.M. of MT-II/GAPDH mRNA ratios.
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Results |
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MT-II mRNA levels in skeletal muscle biopsies increased gradually (P < 0.05) following one bout of exercise, reaching 11- to 15-fold increases at 24 h after exercise, whereas MT-II mRNA was hardly detectable in resting controls (Fig. 1)
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Resting subjects demonstrated very low or zero MT-I + II expression in the muscle tissue (Fig. 2A–C). By the end of the exercise period (3 h), the immunohistochemical staining for MT-I + II was increased in all fibre types of the muscle tissue (Fig. 2D). However, some muscle fibres were strongly immunoreactive for MT-I + II, while others showed a smaller increase. This variety in MT-I + II staining intensity was only pronounced at 3 h when the exercise ended, and it could not be attributed to muscle fibre types, as judged by comparing myofibrillar ATPase staining (not shown) with MT-I + II immunohistochemistry in neighbouring sections. After 4.5 h and especially after 6–9 h, the levels of MT-I + II had further increased (Fig. 2E–G), and by then the tissue showed a very strong, homogenous MT-I + II immunostaining throughout the muscle biopsy. Accordingly, every fibre type of the muscle (Type I, Type IIa or Type 2x) displayed increases in MT-I + II. By 24 h, the levels of MT-I + II proteins had further increased (Fig. 2H), and all muscle fibres showed intense and homogenous MT-I + II immunohistochemical staining. There was no MT-I + II immunostaining present in between muscle fibres. Thus, exercise significantly increased MT-I + II expression levels, which remained highly increased by 24 h post exercise (Fig. 2).
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In order to determine oxidative stress levels, we determined NITT and MDA immunoreactivity.
Muscle tissue from resting subjects showed no immunostaining for NITT (Fig. 3A–C) and MDA, while after the exercise, an increase in NITT (Fig. 3) and MDA (not shown) immunoreactivity was observed. Hence, right after the exercise at 3 h, both NITT (Fig. 3D) and MDA immunoreactivity clearly increased and the levels remained high until 24 h. However, NITT (Fig. 3E) and MDA levels peaked by 4.5 h. Accordingly, the levels of oxidative stress are high, peaking shortly after the exercise and before MT-I + II expression reached high levels.
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Discussion |
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As control participants (the resting subjects) had biopsies taken at identical time points as the subjects who performed exercise, it is unlikely that the oxidative stress observed after 3 h exercise could be caused simply by the recurring biopsies that could induce inflammation and ROS formation by infiltrating leucocytes.
However, the observed dynamics of MT-II mRNA versus MT-I + II protein expression are rather surprising, as the mRNA increases should precede those of the proteins. This might be linked to the fact that the immunohistochemistry is presumably detecting all or most of the many human MT-I + II isoproteins, while the mRNA measured was only for a subtype of MT-II (subtype MT-IIa). As it is not possible to raise an antibody against either the MT-I or the MT-II isoprotein without mutual cross-reaction, this should be addressed by additional detection of the different types of MT-I mRNA as well as MT-II mRNA. On the other hand, the observed levels of mRNA versus proteins could be explained if human MT-I mRNA is induced and translated faster relative to MT-II mRNA. Moreover, it can not be ruled out that muscle MT-II mRNA translation is more efficient during exercise or is enhanced in the first days after exercise. The MT-II mRNA levels observed in the 24 h post-exercise samples showed a high inter-individual variation, and in support of this, MT-I + II display extensive genetic polymorphism (Hidalgo et al. 2001, 2002). Even though, the MT-I + II proteins did not show such variance between the participants.
These issues have to be addressed in the future, and measurements of the different types of MT-I mRNA as well as MT-II mRNA need further attention.
Regardless of the factors involved, this induction of MT-I + II by physical training may be of importance in protection against ROS and oxidative stress in the muscle, as MT-I + II are extraordinarily efficient ROS scavengers and antioxidant proteins (Thornalley & Vasak, 1985; Aschner, 1996, 1998; Molinero et al. 1998; Shinogi et al. 1999; Kumari et al. 2000; Rana & Kumar, 2000; Viarengo et al. 2000; Hidalgo et al. 2001, 2002; Penkowa et al. 2000; Penkowa, 2003; Theocharis et al. 2003). Accordingly, many in vitro studies have shown that MT-I + II inhibit ROS-induced DNA degradation and tissue damage (Schwarz et al. 1994, 1995; Lazo & Pitt, 1995; Lazo et al. 1995; Pitt et al. 1997; Jiang et al. 1998; Haidara et al. 1999; Cai et al. 2000; Kling & Olsson, 2000; Suzuki et al. 2000). In fact, MT-I + II could protect against ROS-induced DNA damage in vitro with much higher molar efficiency (almost 800-fold) when compared with glutathione (Abel & de Ruiter, 1989). Also, MT-I + II can functionally substitute for Cu/Zn-SOD in the cellular defence against oxidative stress (Tamai et al. 1993), and MT-I + II can compensate for Cu/Zn-SOD deficiency in mice (Ghoshal et al. 1999). In vivo, MT-I + II are very powerful antioxidants and scavengers of free radicals, and mice with genetic MT-I + II deficiency (MT-I + II knock-out (MT-I + IIKO) mice) show significantly increased levels of ROS formation and oxidative stress relative to wild-type mice, which is also reflected by increased clinical findings and disease manifestations in the MT-I + IIKO mice (Fu et al. 1998; Carrasco et al. 2000; Penkowa et al. 2000, 2001; Hidalgo et al. 2001, 2002; Giralt et al. 2002a; Trendelenburg et al. 2002). It is noteworthy that in MT-I + IIKO mice the levels of antioxidants catalase, Mn-SOD and Cu/Zn-SOD were increased in response to oxidative stress induced by a head trauma relative to wild-type mice (Penkowa et al. 2000). Thus, when MT-I + II were absent, even an increased expression of other antioxidants could not protect from increased oxidative stress levels, which indicates the protective importance of MT-I + II; while mice with MT-I overexpression (MTTg mice) are protected from oxidative stress and show very low levels of ROS (Fu et al. 1998; Chen et al. 2001; Giralt et al. 2002b; Penkowa et al. 2002; Molinero et al. 2003) relative to wild-type mice. Additionally, MT-II can be used therapeutically for in vivo pathological conditions with oxidative stress (Penkowa & Hidalgo, 2000; Giralt et al. 2002b; Penkowa et al. 2002). Also, MT-I + II were beneficial, as increases in MT-I + II production or transgenic MT-II overexpression in pancreatic islets of mice could prevent diabetes induced with streptozotocin (Ohly et al. 2000; Chen et al. 2001). Thus, transgenic MT-II overexpression could drastically reduce pancreatic islet disruption, cell death, DNA damage and depletion of nicotinamide adenine dinucleotide (NAD+) as well as clinical disease and hyperglycaemia (Chen et al. 2001).
In addition, overexpression of MT reduces diabetic cardiomyopathy effectively (Liang et al. 2002; Ye et al. 2003), and MT could eliminate the increased ROS formation in the myocytes during diabetes (Ye et al. 2003).
From the data presented here we can not determine the relative contribution of the increased MT-I + II to muscle total antioxidant capacity. However, during specific conditions with impaired human MT-I + II signalling, exercise-induced increases in oxidative stress are enhanced (authors' own unpublished data). Therefore it is likely that the observed increase in MT-I + II presented here is physiologically important.
As mentionend in detail above, scientific data suggest that MT-I + II are important for the tissue oxidative balance. Moreover, sections from the present study stained for other antioxidant factors (e.g. Cu/Zn-SOD, Mn-SOD, myeloperoxidase and catalase) show reduced expression levels relative to those of MT-I + II proteins after exercise. Hence, the MT-I + II increase shown here is the most pronounced antioxidant response within the time period studied after exercise. Even if that does not rule out the possibility that another antioxidant may increase even more at later time points, MT-I + II are at least manifest during the peak of exercise-induced oxidative stress.
In addition, many studies have shown that MT-I + II protect against oxidative stresses in various tissues (for review see Viarengo et al. 2000; Hidalgo et al. 2001, 2002). Accordingly, the increased MT-I + II expression observed in muscle tissue after physical exercise probably has important roles in protection against exercise-induced ROS formation and oxidative stress, which indeed were increased in the present study. In addition, the finding that non-damaging exercise induces a strong antioxidative activity may also represent yet another mechanism whereby exercise protects against chronic medical disorders such as atherosclerosis, obesity, cardiovascular diseases and diabetes (Ruderman & Schneider, 1992). In support of this, MT-I + IIKO mice develop obesity including increased obese (ob) gene expression, elevated fat accumulation and high concentrations of plasma leptin, all symptoms that are similar to those recorded in Zucker fatty (fa/fa) rats. Thus, an association between MT-I + II and energy homeostasis has been implied. Thus, mice with MT-I + II deficiency develop spontaneous obesity and hyperleptinaemia (Beattie et al. 1998). In conclusion, the finding that non-damaging exercise markedly increases MT-II mRNA and MT-I + II protein in human skeletal muscle fibres is likely to represent an antioxidant defence system after physical exercise, and thereby MT-I + II induction may contribute to the beneficial metabolic effects of exercise.
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Significant difference between groups. *Significant change over time. MT II mRNA was on the border of detection in the resting control group (n
= 5).
