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Master in sports physiology and medicine
Degree project in sports physiology and medicine
Second level, 45higher education credits
(Spring 2011)
Title:
THE EFFECTS OF MUSCLE DAMAGING ELECTRICALLY STIMULATED
CONTRACTIONS AND IBUPROFEN ON MUSCLE REGENERATION AND
TELOMERE LENGTHS IN HEALTHY SEDENTARY MALES
Author:
Mathias Ekstrand
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ABSTRACT
Introduction: The effect of electrical stimulation on muscle degeneration and regeneration is
largely unknown and it has not been studied in conjunction with telomeres. The consumption of non-steroidal anti-inflammatory drugs (NSAIDs) is widespread in athletes and the general population when faced with muscle soreness or injury. Furthermore, the effect of NSAIDs on muscle regeneration is controversial and its effect on telomere lengths is also unknown.
Methods: Young adult males performed 200 electrically stimulated maximal isokinetic
contractions with one leg (ES) and the other worked as a control (CON). They received either 1200mg ibuprofen (IBU) per day or placebo (PLA) from 21 days pre- to 30 days post-exercise. Muscle biopsies were obtained from the vastus lateralis of the CON leg at baseline (H0) and ES leg at 2.5h (H2.5) and both legs at 2, 7 and 30 days post-exercise. Blood samples were obtained at the same time points and at day 4 post-exercise. Afterwards the muscle and blood specimen were analysed for skeletal muscle and peripheral blood telomere lengths by Southern blot and signs of muscle degeneration and regeneration were quantified histologically. Results: Histological changes occurred in the ES leg, including; increased proportion of damaged myofibres (2.1±2.8%) and infiltrated myofibres (5.0±6.0%) at day 7, small myofibres (3.0±4.4%) and internally located myonuclei (2.9±3.1%) at day 30. The IBU group had significantly less internally located myonuclei at day 30 compared to PLA (1.7±2.4% vs. 4.1±3.8%). No significant differences were observed in skeletal muscle mean and minimum telomere lengths between ES and CON leg, between IBU and PLA group or between time points. Peripheral blood mean telomere lengths were not significantly different between IBU and PLA group, but between time points; H0 (9.6±1.2kb) and H2.5 (9.1±1.1kb) were significantly shorter than day 4 (10.3±1.6kb) and day 7 (10.1±1.5kb) (P<0.05).
Conclusion: Electrically stimulated contractions caused significant muscle degeneration and
regeneration in the 30 days post-exercise. Electrical stimulation also appeared to cause fluctuations in peripheral blood telomere lengths, but not skeletal muscle telomeres. The intake of ibuprofen appeared to interfere with muscle regeneration, but did not seem to affect the peripheral blood or skeletal muscle telomeres. However, due to marked individual variations and the small participant group it is difficult to conclude on the effects of electrical stimulation and ibuprofen on proliferative potential. Further studies are warranted to elucidate the effects of electrical stimulation and ibuprofen on blood and skeletal muscle telomeres.
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INTRODUCTION
Skeletal muscle repair is a highly synchronised process involving the activation of various cellular responses. The first phase of muscle repair is characterised by degeneration of damaged myofibres and activation of an inflammatory response. This phase is followed by the formation of new myofibres through myogenesis (Huard et al., 2002). Because the nuclei within myofibres are postmitotic, myogenesis primarily depends on satellite cells, which are quiescent muscle precursor cells. In response to injury, satellite cells re-enter the cell cycle and proliferate. The satellite cells then undergo differentiation and fusion to restore normal muscle structure. However, human diploid cells have a limited proliferative capacity, as demonstrated on fibroblasts (Hayflick, 1965) and satellite cells in vitro (Decary et al., 1996). The proliferative capacity of human diploid cells is partially limited by the shortening of telomeric sequences (Allsopp et al., 1996).
Telomeres are non-codant repetitive sequences (TTAGGG)n of DNA located at the
end of chromosomes (Blackburn, 1990). Telomeres protect chromosomes from degradation, end-to-end fusion and atypical recombination (Moon and Jarstfer, 2007; Orr-Weaver et al., 1981; Haber and Thorburn, 1984). In healthy humans the telomere sequences are 3-20kb in length (de Lange et al., 1990; Klapper et al., 2001), and shorten by 50-200bp with each cell division, in part due to ineffective replication of the 5’ end by DNA polymerases (Collins and Mitchell, 2002; Olovnikov, 1973). Once the telomeric DNA is too short it becomes senescent and stop dividing, limiting the regenerative capacity of the cell (Decary et al., 1996). Since proliferating satellite cells then lose a part of the telomeric DNA during each cell cycle, the newly replicated DNA in the regenerated myofibre would have a shorter telomere sequence (Decary et al., 1997).Telomere shortening is however not an irreversible process, but can be elongated by a nucleoprotein complex called telomerase (Greider, 1996). Telomerase has been demonstrated to reset the mitotic clock at each generation in the germline in vitro (Rubin, 2002) and telomerase activity has been reported in many normal tissues, including skeletal muscle and blood (Radak et al., 2001; Engelhardt et al., 1997). The number of cycles of satellite cell proliferation along with their remaining regenerative capacity can be determined by measuring the telomere lengths in skeletal muscle biopsies (Kadi et al., 2008; Rae et al., 2010).
When satellite cells are heavily recruited for muscle regeneration as in the skeletal muscle of training athletes, telomere lengths has been found to have a tendency to be significantly longer (Kadi et al., 2008; Kadi and Ponsot, 2010), equivalent (Rae et al., 2010), and, in extreme cases, significantly shorter (Collins et al., 2003) than control subjects.
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Significant increases in the proportion of satellite cells have been detected 4 and 8 days following a single bout of maximal exercise (Crameri et al., 2004) and more than twofold increase in satellite cell number has been demonstrated 24h after maximal eccentric contractions in young males (Dreyer et al., 2006). The largest acute increase in satellite cell number has been observed following voluntary or electrically induced maximal eccentric contractions (Crameri et al., 2004, 2007; Dreyer et al., 2006). The significant recruitment of satellite cells in the adaptive process of skeletal muscle to exercise gives rise to the question of whether an acute bout of electrically stimulated muscle contractions affects skeletal muscle DNA telomere length.
Measuring telomere lengths in the peripheral blood present a promising new perspective in understanding the peripheral effects of strenuous exercise. Peripheral blood cell telomere lengths correlate well with hematopoietic stem cells (Sakoff et al., 2002), which are pluripotent cells that give rise to all blood cell types (Abramson et al., 1977). A multitude of hematopoietic stem cell subpopulations (from myeloid macrophages and neutrophils to lymphoid T- and B-cells) are key players in orchestrating the body’s response to environmental stress, such as strenuous exercise (Fitzgerald, 1988; Kierszenbaum, 2007; Tidball and Vilalta, 2010). No study has to our knowledge investigated the effects of muscle damaging exercise on peripheral blood telomere lengths.
The most commonly used model for studying exercise-induced muscle damage and subsequent regeneration is one involving voluntary eccentric contractions. However, recent evidence suggests that the use of electrically stimulated contractions induce even greater muscle damage. Crameri et al.(2007) found disruption of cytoskeletal proteins, destroyed z-lines and the satellite cell response to be markedly more pronounced in the electrically stimulated leg compared to the voluntarily contracted leg after 210 maximal eccentric contractions. Moreover, Mackey et al. (2008) demonstrated direct evidence of damage at the myofibre and sarcomere levels following a stimulated isometric contraction protocol, through the presence of macrophages in myofibre cytoplasm and the disruption of z-lines. Given the multitude of applications of neuromuscular electrical stimulation, from the enhancement of exercise performance in athletes, to prevention of atrophy in patients and elderly, deeper knowledge regarding the physiological adaptations to stimulated contractions is called for. Exercise-induced muscle damage and the resulting muscle soreness, inflammation and dysfunction is often treated with non-steroidal anti-inflammatory drugs (NSAIDs) (Mahler, 2001; Warner et al., 2002). It is estimated that a total of 30 million people worldwide use NSAIDs on a daily basis (Alaranta et al., 2008). Although plenty of research exists on the
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efficacy of NSAIDs, their effects on the inflammatory response and muscle regeneration are not fully understood. Studies have both found the inflammatory cell response to be inhibited and to be unaffected by NSAIDs ingestion following muscle damaging exercise or injury (Peterson et al., 2003; Bondesen et al., 2004). Furthermore, NSAIDs may negatively affect the satellite cell response to exercise in humans (Mackey et al., 2007; Mikkelsen et al., 2009; Shen et al., 2005; Zalin, 1977, 1987). Mikkelsen et al. (2009) found a 96% increase in satellite cell count following maximal eccentric contractions. NSAIDs abolished this increase. The effect of NSAIDs on telomere length is presently unknown.
In the present study we investigated the effects of electrically stimulated eccentric contractions on muscle damage, regeneration, and telomere length in both peripheral blood and skeletal muscle. We also investigated the influence of NSAIDs ingestion on muscle degeneration, regeneration and telomere length.
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METHODOLOGY
Participants
Thirty-three participants (age 21 ± 2.72 years, height 180.7 ± 6.19cm and body mass 73.3 ± 7.96kg) were recruited for the study after being interviewed and screened for kidney and liver dysfunction. Four were excluded from the histological analysis as some of their histology slides were missing or of insufficient quality. Peripheral blood and skeletal muscle telomere lengths were supposed to be assessed in 14 and 12 subjects, respectively. However, due to experimental anomalies and insufficient material, 11 subjects were assessed for peripheral blood telomere lengths and 9 subjects for skeletal muscle telomere lengths in this study. All participants were unaccustomed to high-intensity exercise and were not taking part in any regular exercise. The Ethics Committee of the Municipalities of Copenhagen and Frederiksberg approved this study, and all procedures conformed to the Declaration of Helsinki. This study was performed in collaboration with the institute of Sports Medicine in Copenhagen.
Study Design
2 weeks pre-test. The participants were assigned to two groups matched for age, height and
body mass. The first group (IBU) ingested 600mg ibuprofen (Ibumetin, Nycomed Denmark Aps) twice per day from 14 days pre-exercise, until 30 days post-exercise. The second group (PLA) ingested placebo in the same timeframe. The study was double-blind. Blood samples were taken to ensure that the participants were complying with the agreed drug intake, and ingested no other anti-inflammatory substances.
Exercise day. Blood was sampled and a muscle biopsy (baseline) was obtained from the
non-exercising leg (CON) pre-exercise. After a minimum of 1h of rest, participants performed 200 maximal electrically stimulated eccentric contractions in an isokinetic dynamometer. Blood was sampled and a muscle biopsy was obtained from the exercising leg 2.5h post-exercise (H2.5).
Day 2 to day 30 post-exercise. Biopsies were obtained from the electrically stimulated leg (ES) and the CON leg at 2 (D2), 7 (D7) and 30 days (D30) post-exercise. Blood was sampled at D2, day 4 (D4), D7 and D30 post-exercise.
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Exercise Protocol
The exercise protocol was adapted from Crameri et al. (2007). One leg was randomly assigned to perform exercise using percutaneous electrical stimulation, the other leg worked as a control (CON). Participants were to sit in an isokinetic dynamometer, with their backs at a 10˚ angle (KinCom, Kinetics Communicator, Farrell and Richards, 1986). The rotational axis of the dynamometer lever arm was set parallel with the lateral femoral epicondyle. The lower lever arm was attached to the lower leg approximately 2cm above the lateral malleolus, without static fixation of the ankle joint (Aagaard et al., 2000). As described by Aagaard et al. (1995), the force signal was corrected for the effect of gravity at an angle of 45˚. Two electrodes (Bio-Flex, 50mm x 89mm, Biofina A/S, Odense, Denmark) were attached to the vastus lateralis muscle of the exercise-leg. The first was attached 5cm from the distal end of the patella and the second 5cm from the anterior superior iliac spine (Theriault et al., 1996). Impulse trains (300µs single pulse duration; 35Hz; maximal current, 100mA) were delivered to the vastus lateralis muscle using a constant-bicurrent stimulator (ELPHA II 2000, Biofina, Odense, Denmark, Crameri et al., 2004). The participants were familiarised to the electrical stimulation by gradual increase of the current until they reached their highest tolerable limit. The protocol consisted of two phases: (1) 100 maximal quadriceps contractions (5 sets of 20 repetitions) at slow contraction speed (30˚ s-1 knee joint angular velocity); followed by (2) 100 maximal quadriceps contractions (5 sets of 20 repetitions) at high contraction speed (180˚ s-1). Downward movement of the leg by electrical stimulation commenced at 10˚ knee joint angle and ended at 90˚, after which the leg immediately returned to the starting position by means of the motor of the dynamometer. The participants were carefully instructed to not produce any voluntary muscle force of the stimulated-leg during the exercise protocol. Between each set the participants rested for 30s and for 5min between the slow and fast contraction phases.
Isolation of peripheral blood cells
Four ml of blood were collected in BD Vacutainer® CPT™ Tube from the antecubital vein in a supine position. Mononuclear cells were then isolated from whole blood according to manufacturer's instructions, frozen and stored at -80°C.
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Muscle Biopsies
Percutaneous needle biopsies were performed under local anaesthesia (1% lidocaine). Muscle samples were collected from the vastus lateralis muscle using a 5-mm Bergström biopsy needle (Evans et al., 1982). The muscle biopsies were quickly divided for histological and biochemical analysis, frozen in liquid nitrogen and stored at -80°C.
Histological Analysis of Muscle Degeneration and Regeneration
The muscle specimens used for histological analysis were mounted cross-sectionally on cork with Tissue-Tek OCT mounting medium and sectioned 8µm thick in a cryostat microtome (Microm, Cryo-Star HM 560M) cooled to -20°C. The specimens were then mounted on glass coverslips, air dried, and stained with haematoxylin and eosin. Images of the sections were obtained using a digital camera (SPOT Insight, Diagnostic Instruments) connected to a Nikon microscope (Eclipse E400). Damaged muscle fibres (fibres showing loss of normal polygonal outline, hyaline aspect and/or disruption of the sarcolemma), fibres infiltrated by mononuclear cells, fibres with internal nuclei and small muscle fibres were quantified and divided by the number of muscle fibres in the specimen and multiplied by a 100 to get the percentage (Lexell et al., 1992; Rosenblatt and Woods, 1992; Van der Meulen, 1991). A test-retest analysis was done to estimate the reliability with which each of the numerical ratio could be made. For all variables, the coefficient of variation was less than 3%. The muscle fibres were counted in MuscleFibre Deluxe v1.0.
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Figure 1: Representative histological images of the four parameters used to quantify muscle degeneration and regeneration as seen in the stimulated leg. A) Damaged (white arrows) and infiltrated muscle fibres (red arrows) B) Small muscle fibres (orange arrows) and internally located muscle fibres (blue arrows).
Telomere Length Analysis
Mean and minimum telomere lengths were assessed using a Southern blot protocol (Collins et al., 2003; Decary et al., 1997; Renault et al., 2002). The protocol was modified for the study of human skeletal muscle and blood, as delineated by Ponsot and Kadi (2007). Muscle and blood nuclei were obtained after separation of cytoplasm and nuclear using the NE-PERTM Nuclear and Cytoplasmic Extraction Reagents Kit (Pierce Biotechnology, Rockford, USA). For the extraction of genomic DNA,the nuclei was digested overnight in lysis buffer (Triton, 10mM Tris-Cl, pH 8.0, 1mM EDTA, pH 7.5, NaCl, 0.5M) and 300µg/ml Proteinase K at a temperature of 55˚C. Nucleic acids were extracted with Phenol/chloroform/isoamyl alcohol (25: 24: 1 (vol:vol:vol)). DNA was precipitated with 0.5 vol of ammonium acetate 7.5M, and 2 vol of ethanol 100%. After 48h precipitation in -20˚C, the DNA was washed in ethanol 70%, resuspended and stored at 4˚C in TE buffer (10mM Tris-Cl, pH 8.0, 1mM EDTA, pH 7.5). The genomic DNA was digested for 5h at 37˚C with the restriction enzyme
Hinf I (Biolabs, New England). This enzyme produces telomere restriction fragments with
different lengths of the TTAGGG repeat sequence and a constant sub-telomeric fragment (Decary et al., 1997). Three micrograms of digested genomic DNA, together with a high molecular weight and 1 kb DNA ladder were resolved for 20h by electrophoresis (90V) in
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+4˚C on 0.7% agarose gels (figure 2). Gels were dried, denatured and then neutralised. The telomere restriction fragments were detected directly in the denatured and neutralised gels by hybridisation to a 32P-labeled (TTAGGG)4 probe, then exposed to a film. The signal
responses were scanned, analysed in Scion Image and then modelized as described by Ponsot and Kadi (2008).
Figure 2: Representative autoradiograms showing the telomeric DNA from peripheral blood (left) and skeletal muscle (right) after 20h migration at 90V in an 0.7% agarose gel.
Statistical Analysis
All statistical analysis was performed in SigmaPlot v11.0 and Excel 2010. Where applicable the data is presented as mean ± standard deviation (SD) and the significance level is set to 5%. Two-way repeated measures ANOVA with holm-sidak post-hoc were used to test significant difference between ES and CON leg, between the treatment groups and time points. To determine any potential correlations between variables Pearson Product Moment Correlation tests were conducted. There was only enough data at H0, H2.5 and D30 to perform statistical analysis on skeletal muscle telomere lengths between the treatment groups (figure 6). When comparing skeletal muscle telomere lengths in ES and CON leg, the treatment groups are combined into one larger group (figure 5). This is also the case when comparing peripheral blood telomere lengths between different time points (figure 7b).
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RESULTS
Morphology
Analysing the electrically stimulated leg in both the NSAIDs and placebo group, the proportion of damaged (2.1±2.8%) and infiltrated muscle fibres (5.0±6.0%) was significantly elevated at day 7 compared to all other time points and compared to control leg (0±0% and 0±0%, respectively, P<0.05, figure 3a & b). In the ES leg the proportion of damaged muscle fibres ranged between 0 and 12.2%, and 0 and 8.2% in the NSAIDs and placebo group, respectively (figure 4). The proportion of infiltrated muscle fibres ranged between 0 and 24.7%, and 0 and 15.8% in the ES leg in NSAIDs and placebo group, respectively. The proportion of small muscle fibres (3.0±4.4%) and muscle fibres with internal nuclei (2.9±3.1%) was significantly elevated at day 30 compared to all other time points and compared to control leg, in both treatment groups (0±0% and 0±0%, respectively, P<0.05, figure 3c & d). The IBU group had significantly less internally located myonuclei in the ES leg at day 30 compared to PLA (1.7±2.4% vs. 4.1±3.8%, P<0.05, figure 3d). No significant differences were detected in the control leg in either group, nor between groups for any parameter.
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Figure 3: The proportion of degenerating (A, B) and regenerating muscle fibres (C, D) at the different time points in the electrically stimulated leg (n=29). * P<0.05 vs. all other time points; #: P<0.05 vs. other treatment group.
Figure 4: Individual profiles for the proportion of damaged and infiltrated muscle fibres in (A) the NSAIDs group and (B) the placebo group at day 7.
0 2 4 6 H2.5 D2 D7 D30 % Dam ag e d M u scle Fi b re
s
Damaged Muscle Fibres
IBU PLA
A
* * 0 2 4 6 8 10 12 H2.5 D2 D7 D30 % In fi litr ated M u scl e Fi b resInfiltrated Muscle Fibres
IBU PLA
B
* * 0 2 4 6 8 10 H2.5 D2 D7 D30 % Sm al l M u scle Fi b re sSmall Muscle Fibres
IBU PLA
C
* * 0 2 4 6 8 10 H2.5 D2 D7 D30 % In te rn al N u cl e iInternal Nuclei
IBU PLAD
* * # 0 5 10 15 20 25 30 S1 S3 S6 S8 S12 S13 S15 S17 S20 S23 S24 S27 S29 S31 Pr o p o rtion D 7 ( % ) SubjectsNSAIDs
dmg fibresinfiltratedA
0 5 10 15 20 S5 S9 S10 S11 S14 S18 S19 S21 S22 S25 S26 S28 S30 S32 Pr o p o rtion D 7 ( % ) SubjectsPlacebo
dmg fibres infiltratedB
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Telomere Lengths
Irrespective of treatment group, skeletal muscle mean telomere lengths (Lmean) and minimum
telomere lengths (Lmini) were not significantly different in the ES leg compared to CON leg
for any of the time points (Figure 5).
Figure 5: Skeletal muscle (A) Lmean and (B) Lmini in the electrically stimulated leg (ES) and
control leg (CON) irrespective of treatment group (n=18).
Skeletal muscle Lmean and Lmini were not significantly different at any time point between the
NSAIDs and placebo group (figure 6).
Figure 6: Comparison of skeletal muscle (A) Lmean and (B) Lmini inthe electrically stimulated leg
between the NSAIDs and placebo group (n=9). H0 correspond to the control leg. 0 2 4 6 8 10 12 14 16 D0 D2 D7 D30 M e an Tel o m e re Le n gth ( kb ) CON ES
A
H0 H2.5 0 1 2 3 4 5 6 7 D0 D2 D7 D30 M in im u m Te lo m e re Len gth ( kb ) CON ESB
H0 H2.5 0 2 4 6 8 10 12 14 16 D0 H2.5 D30 M e an Tel o m e re Le n gth ( kb ) NSAIDs PLACEBOA
H0 0 1 2 3 4 5 6 7 D0 H2.5 D30 M in im u m Tel o m e re Len gth ( kb ) NSAIDs PLACEBOB
H014
Peripheral blood Lmean were not significantly different between the NSAIDs and placebo
group (figure 7a). Significant differences existed between several time points within each treatment group in peripheral blood Lmean (P<0.05, figure 7a). Irrespective of treatment group,
the peripheral blood Lmean were significantly shorter at H0 (9.6±1.2kb) and H2.5 (9.1±1.1kb)
compared to D4 (10.3±1.6kb) and D7 (10.1±1.5kb) (P<0.05, figure 7b).
Figure 7: (A) mean peripheral blood telomere lengths in the NSAIDs and placebo groups (n=11). (B) mean peripheral blood telomere lengths irrespective of treatment group (n=11). * P<0.05 vs. D4; #: P<0.05 vs. D7
Skeletal muscle Lmini were significantly negatively correlated with the proportion of
infiltrated myofibres at day 2 (r = -0.81, P<0.05) and day 7 (r = 0.77, P<0.05). 0 2 4 6 8 10 12 14 H0 H2.5 D2 D4 D7 D30 M e an Tel o m e re Le n gth ( kb ) NSAIDs PLACEBO
A
* # * * 0 2 4 6 8 10 12 14 H0 H2.5 D2 D4 D7 D30 M e an Tel o m e re len gth s (k b )B
*
# #*
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DISCUSSION
Morphology
The electrical stimulation protocol caused significant damage and inflammation in the muscle fibres of the stimulated leg at day 7, as evidenced by histological quantification of degenerating myofibres (showing loss of normal polygonal outline, hyaline aspect and/or disruption of the sarcolemma, and infiltration of mononuclear cells). Recently, Tidball and Vilalta (2010) outlined a timeline of the inflammatory cell response following acute muscle damage. Neutrophils are the first inflammatory cells to respond to muscle injury. They begin to invade the site of injury within 2h of muscle damage, typically peak in concentration between 6 and 24h post-injury, and then quickly diminish in numbers (Fielding et al., 1993; Quindry et al., 2003). At 24h post-injury, M1 macrophages reach significant numbers at the site of injury. These phagocytic M1 macrophages continue to increase in numbers until 2 days post-injury, when they rapidly decline (Ochoa et al., 2007). An invasion of non-phagocytic M2 macrophages ensues, peaks at approximately 4 days post-injury and remains at the site of injury for many days (Tidball and Villalta, 2010). Therefore, the presence of mononuclear cells in the myofibres at day 7 likely represents the invasion of M2 macrophages. Other studies have demonstrated similar muscle damage and inflammation following electrical stimulation. Using an identical electrical stimulation protocol to ours, Crameri et al. (2007) found a significant increase in desmin-negative myofibres, myofibre infiltration by macrophages (unknown phenotype), and destroyed and disrupted z-lines in human vastus lateralis muscle at time points between 1 and 8 days post-exercise compared to baseline. Similarly, Mackey et al. (2008) demonstrated significant myofibre infiltration by macrophages (CD68+ M1 macrophages), loss in desmin-negative myofibres and disrupted z-lines in human gastrocnemius medialis muscle two days after an extensive electrically stimulated isometric muscle contraction protocol. However, two days after electrical stimulation we found only modest amounts of infiltrated muscle fibres (five out of 29 subjects had infilitration of 0.7–1.4 %, with the rest failing to show any infiltration). Yet, examining individual profiles in the study by Mackey and colleagues, the proportion of macrophage-infiltrated myofibres ranged from 0.1 to 0.7% in four out of seven participants. One individual responded with a markedly elevated infiltration of 12.7% and two failed to show any macrophage infiltration. These results are rather similar to our results at day 2, if one excludes the outlier. Likewise, our results showed remarkable individual variation in the proportion of infiltrated myofibres at day 7 as shown in figure 4. It is possible that Mackey and colleagues also would have observed more inflammation and muscle damage at later
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time points. What is more in the previously mentioned study, Mackey and colleagues demonstrated that the extent of muscle damage is related to the stimulated force produced by the muscle, suggesting that force is responsible for myofibre damage. Although the force production was not measured in our study, a similar trend is likely present, suggesting that the electrical stimulation was insufficient to cause muscle damage in some individuals, hence the large inter-individual variations. However, one needs to have in mind when analysing cross sections of muscle biopsies that such a small sample is used to estimate damage in an entire muscle. Moreover, because the damage may not evenly spread throughout a muscle, but rather is focalized, it is possible to over- or underestimate the damage (Clarkson and Hubal, 2002).
Histological analysis also revealed significantly increased proportions of internal nuclei and small muscle fibres in the electrically stimulated leg at day 30 post-exercise. Small muscle fibres and myofibres with a central location of the myonuclei are both commonly regarded as indicative of myofibre regeneration. The central myonuclei are believed to be the progeny of myoblasts which have fused into myotubes (McGeachie et al., 1993). As the myotubes mature, they retain their nuclei in the centre of the fibre for many months after being formed (McGeachie et al., 1993). Hence, the presence of centrally located nuclei in muscle fibres indicated that a cycle of regeneration has occurred. Expectedly, inter-individual variation also occurred in the proportion of small muscle fibres and central myonuclei, with the subjects demonstrating myofibre damage and infiltration also showing signs of regeneration, emphasizing the link between muscle degeneration and subsequent muscle regeneration. In accordance with our results, studies have demonstrated macrophages to aid in muscle injury and clear evidence show macrophage involvement in muscle regeneration and growth following muscle injury (Chazaud et al., 2003; Tidball and Vilalta, 2010).
The proportions of damaged and infiltrated muscle fibres were not significantly different between the ibuprofen and placebo groups in the electrically stimulated leg. Therefore, our results suggest that ibuprofen does not affect myofibre inflammation and damage. Several animal studies have reported reduced levels of inflammation both in blood and muscle with NSAIDs ingestion following extensive exercise and injury (Bondesen et al., 2004; Novak et al., 2009; Shen et al., 2005, 2006), whereas the effects of NSAIDs on the inflammatory response in muscle and blood is equivocal in humans Mikkelsen et al., 2009; Peterson et al., 2003; Weinheimer et al., 2007). Peterson et al. (2003) administered the same
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dose of ibuprofen (1200mg/day) as in the current study, acetaminophen (4000mg/day) or placebo to males after maximal eccentric contractions of the knee extensors. The macrophage concentration in the blood increased 1.5-2.5fold at 24h post- compare to pre-exercise, and as in our study, were not significantly different between treatment groups and placebo. Mikkelsen et al. (2009) used the exact same exercise protocol as was used in the current study, but with voluntary exercise contractions instead of electrically stimulated ones. In accordance with the conclusions of the study of Crameri et al. (2007), they found neither significant increase in desmin-negative fibres, nor infiltration of inflammatory cells into the myofibres 8 days post-exercise. As in our study, large individual variations in the inflammatory response were reported in the study by Mikkelsen et al., with some individuals demonstrating 2-6fold increase compared to pre-exercise. Furthermore, they infused indomethacin via a microdialysis catheter into the vastus lateralis muscle of one leg immediately before and up until 4.5 h after the 200 maximal eccentric contractions. No significant differences in inflammation were detected between the NSAIDs leg and the control leg. The exercise stimuli using voluntary eccentric contractions were likely insufficient to cause damage to the myofibres, emphasizing together with our results, and the results of Crameri and colleagues the increased muscle damage caused by electrical stimulation. NSAIDs are known to block the COX pathways that contribute to cell-mediated prostaglandin production (Weinheimer et al., 2007). The role of prostaglandins in muscle inflammation can be challenging to untangle in vivo because they are known to exert pro- (PGE2) and anti-inflammatory (PGF2α) properties (Gilroy et al., 1999; Willoughby et al.,
2000) and because of a host of prostaglandin-independent factors that may influence the trafficking of inflammatory cells into damaged muscle (Grounds and Davies, 1996). While the likely function of ibuprofen is in the prevention of prostaglandin synthesis through COX inhibition, it has also been suggested that ibuprofen can act through various COX-independent mechanisms, inhibiting some pathways and activating others (Tegeder et al., 2001). Hence, the lack of treatment effect on inflammation in this study may reflect the inability of therapeutic doses of ibuprofen to inhibit mediators of inflammation, while attenuating modulators of inflammation (Willoughby et al., 2000). Furthermore, COX-2 has been shown to affect muscle cells directly. In a study by Otis et al. (2005) cyclical stretch of cultured myoblasts caused an increase in COX-2 expression and inhibition of COX-2 abolished the stretch-induced myoblast proliferation. This suggests that NSAIDs can inhibit muscle regeneration without inhibiting inflammatory cell invasion by acting directly on the muscle cells.
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The proportion of myofibre with internal nuclei was significantly reduced in the ibuprofen group compared to placebo at day 30 post-exercise in our study, whereas the difference in the proportion of small muscle fibres was non-significant. While it can take months for centrally located myonuclei to migrate to the fibre’s periphery (McGeachie et al., 1993), at 30 days post-exercise newly formed myofibres may have grown to such a size that they are no longer distinguishable from other myofibres (Chargé and Rudnicki, 2004). This possibly explains the lack of significant difference in the proportion of small muscle fibres between the groups. Anti-inflammatory drugs have been shown to inhibit satellite cell proliferation (Shen et al., 2005; Zalin, 1987), differentiation (Schutzle et al., 1984; Shen et al., 2005) and fusion in vitro (Zalin, 1977). Prostaglandins have also been shown to aid in the regulation of muscle fibre growth and muscle protein synthesis and degradation (Trappe et al., 2001). Bondesen et al. (2004) found that SC-236, which is a COX-2-specific inhibitor, decreased the size of regenerating myofibres in rats 5 weeks after freeze injury. Just like in our study they had ongoing administration of the anti-inflammatory drug until the end of the experiment. The attenuation of myofibre growth was associated with decreases in the number of myoblasts and intramuscular inflammatory cells. Interestingly, SC-236 had no effect on myofibre growth when administered starting 7 days after injury. Unlike Bondesen and colleagues, we found no decrease in inflammatory cell response with NSAIDs ingestion in our study, but this discrepancy between studies can be attributed to several factors, including differences in species, drug type and dosing, type of muscle injury and quantification of inflammatory response. Interpretation of our results in combination with the previously mentioned studies may indicate that in addition to being an ineffective treatment for inflammation, therapeutic doses of ibuprofen may negatively regulate muscle regeneration following electrical stimulation.
Telomere Lengths
The skeletal muscle Lmean and Lmini were not significantly different between the ES leg and
the CON leg. The skeletal muscle Lmean and Lmini in both ES and CON legs were within the
range of values previously recorded in skeletal muscle DNA in healthy populations (Decary et al., 1997; Kadi et al., 2008; Ponsot et al., 2008; Renault et al., 2002). Lmean is often used to
examine telomere length in many tissues, both in studies of healthy and unhealthy individuals. However, in skeletal muscle which is a post-mitotic tissue, it has been
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demonstrated that Lmean is a less suitable measure of the relatively small loss of telomeric
DNA that occurs in these cells (Kadi et al., 2008). Lmean comprises telomeres of the most
post-mitotic myonuclei which had been incorporated into the muscle fibres from birth, and therefore have been subjected to few mitotic divisions prior to differentiation. On the other hand, the Lmini corresponds to telomeres from satellite cells and myonuclei incorporated
during the last cell divisions (Decary et al., 1997; Ponsot and Kadi, 2008). Hence, the Lmini
better reflects changes in muscle turnover, which is crucial when analysing the regenerative potential of skeletal muscles. Nevertheless, one bout of severe muscle damaging exercise by electrical stimulation was not sufficient to induce significant changes neither in skeletal muscle Lmean, nor Lmini. This may suggest that the satellite cells do not go through frequent
enough rounds of cell division to significantly induce telomere shortening following one bout of severe exercise. It may also suggest that the telomerase activity offset the telomere shortening. Since, no one has previously investigated the acute effects of exercise on telomere length, nor has anyone taken repeated measurements of skeletal muscle telomere length in the same individual before, this is difficult to comment on. Interestingly, the skeletal muscle Lmini in the electrically stimulated leg were significantly negatively correlated to the
proportion of infiltrated myofibres at day 2 and day 7. The negative correlation possibly means that the more myofibres that are infiltrated by mononuclear cells, the greater the proliferation of satellite cells. Indeed, TNFα released by mononuclear cells (neutrophils and M1 macrophages) at the site of muscle damage following injury have been shown to attract satellite cells to the site of injury, thereby promoting muscle regeneration (Tidball and Vilalta, 2010; Torrente et al., 2003). However, one cannot eliminate the possibility that the nuclear fraction from which the skeletal muscle telomeres are extracted can contain nuclei from other cell types, which might have different telomere lengths to skeletal muscle telomeres.
The peripheral blood Lmean fluctuated significantly between time points. Firstly,
blood cells have a very high turnover. Telomere length in hematopoietic stem cells can be determined from that of peripheral blood cells (Sakoff et al., 2002). Hematopoietic stem cells are pluripotent cells that give rise to all blood cell types from the myeloid (neutrophils, monocytes and macrophages, basophils, eosinophils, erythrocytes, platelets, dendritic cells) and lymphoid lineages (T-cells, B-cells, NK-cells) (Abramson et al., 1977). Only 0.01% of blood cells are hematopoietic stem cells, but they are responsible for the daily production of one trillion blood cells (Ogawa, 1993). It has been calculated that about 8% of hematopoietic
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stem cells enter the cell cycle every day, and 99% over a 2 month period (Cheshier et al., 1999). Hence, the turnover of blood cells is much greater than that of myonuclei, which have an estimated turnover of 1-2% per week. However, it needs to be stated that the myonuclei turnover was estimated in rats and may not be comparable to humans. Secondly, because of the high turnover the blood cells are easily affected by the environment (Lansdorp, 2007). Changes have been shown to occur to a myriad of blood cell types following strenuous exercise (Fitzgerald, 1988). Both lymphoid and myeloid cells are present at the site of insult within hours (Kierszenbaum, 2007; Tidball and Vilalta, 2010). Therefore, it is possible that fluctuations in peripheral blood Lmean were caused by the electrically stimulated contractions,
but one cannot rule out that it may have been caused by other environmental factors, such as stress (Qureshi et al., 2002). Thirdly, the possible involvement of telomerase-dependent regulation pathways cannot be excluded. In an in vitro study, within 48 to 72 hours of culture of CD34+ cells (hematopoietic stem cells) in the presence of cytokines, IL-6, erythropoietin and granulocyte colony-stimulating factor, telomerase activity was shown to be up regulated. The telomerase activity peaked after 7 days in culture and decreased either to baseline or below detection levels within three or four weeks (Engelhardt et al., 1997). The peripheral blood Lmean follow a similar pattern to that of the telomerase activity in vitro, with a peak in
Lmean 7 days post-exercise, and subsequently returning to baseline length. Hypothetically, an
increased level of telomerase activity could explain the elongation of Lmean at day 4 and day
7.
No significant differences were observed in skeletal muscle or peripheral blood telomere lengths between the NSAIDs and placebo group. While some authors have found no effect of anti-inflammatory drugs on muscle regeneration or satellite cells activity (Thorson et al., 1998), several studies have reported a negative effect of anti-inflammatory drugs on various stages of myogenesis and regeneration (Almekinders and Gilbert, 1986; Bondesen et al., 2004; Mackey et al., 2007; Mendias et al., 2004; Mikkelsen et al., 2009; Mishra et al., 1995; Trappe et al., 2001; 2002). For instance, in a previously mentioned study by Mikkelsen et al (2009) it was shown that following 200 maximal eccentric contractions, infusion of NSAIDs into one of the legs abolished the increase in satellite cell count, while the control leg increased satellite cell count by 96% on day 8 post-exercise. It may be that the lack of difference seen in telomere lengths between the treatment groups suggest that proliferative potential is unaffected by ibuprofen ingestion. However, the small subject number when comparing treatment groups (n=4 in skeletal muscle and n=6 in blood), coupled with marked
21
individual variation in telomere length, render the results open to question. The inter-individual variability of in vivo telomere length in skeletal muscle and peripheral blood has been noted in several other studies (Decary et al., 1997; Kadi et al., 2008; Ponsot and Kadi, 2008; Ponsot et al., 2008; Sakoff et al., 2002). Further studies are clearly required to elucidate the effects of NSAIDs on skeletal muscles’ proliferative potential.
Limitations
Several limitations of the present work should be emphasized. First, it is unknown whether the electrical stimulation protocol affected the whole of vastus lateralis muscle or just the superficial myofibres. Electrical stimulation is known to sometimes have limited spatial recruitment of muscle fibres (Vanderthommen et al., 2003). Second, it was unknown whether the NSAIDs intake per se (regardless of electrical stimulation) had any effect on peripheral blood telomere lengths, because the time points prior to ingestion could not be taken into account. Third, in southern blotting, subtelomeric DNA may mask the real telomeric length. Last, because of the great variability of telomere lengths, the small number of subjects per group could not provide definite answers.
CONCLUSION
In conclusion, electrical stimulation caused significant muscle degeneration and regeneration as evidenced by the occurrence of damaged muscle fibres and infiltrated muscle fibres, internal myonuclei and small muscle fibres in the 30 days post-exercise. Electrical stimulation also appeared to cause fluctuations in peripheral blood telomere lengths, but did not seem to affect skeletal muscle telomeres. The intake of ibuprofen appeared to interfere with muscle regeneration as evidenced by a decreased proportion of internally located myonuclei in the ibuprofen group compared to placebo. Ibuprofen did not appear to affect telomere lengths in peripheral blood or skeletal muscle. However, due to marked individual variations and the small participant group it is difficult to conclude on the effects of electrical stimulation and ibuprofen on proliferative potential. Further studies are warranted to elucidate the effects of electrical stimulation and ibuprofen on peripheral blood and skeletal muscle telomeres.
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