Amyotrophic lateral sclerosis (ALS) is an incurable, chronic neurodegenerative disease characterized by selective death of motoneurons in the motor cortex, brainstem, and spinal cord, which controls muscle action [1
]. This disease is phenotypically characterized by loss of muscle tone, paresis, muscle atrophy, and spasticity [2
]. These degenerative changes result in the patient’s loss of overall ability to initiate and control voluntary movements, with the exception of the eyes. Approximately 90% of ALS is a sporadic form of disease (sALS), for which the etiology is unknown, and the remaining percentage is a genetically determined form (fALS). In clinical terms, both forms are virtually identical. Since 1993, the study of the etiology of fALS has focused on mutations in the gene encoding superoxide dismutase type 1 (SOD1), which occurs in nearly one-fifth of all cases of familial form of the disease [3
]. The discovery of the role of mutations in the SOD1
gene in the etiology of ALS has contributed to constructing the most suitable animal test model of the disease (type fALS), containing the human SOD1 G93A
transgene described by Gurney [5
], where glycine is changed to alanine at position 93. The similarity in the course of ALS between the human and animal form allows the study of the pathomechanisms of the disease and testing the factors that may mitigate its course, or even extend the life span, in ALS animals. Notably, causes of neurodegeneration in ALS occur outside of the nervous system. Overexpression of hSOD1 G93A in skeletal muscle not only initiates motoneuron death [6
], but also causes profound muscle atrophy [6
]. Additional support for the view that muscle plays a key role in ALS pathogenesis has been provided by the observations that the destruction of neuromuscular junctions is linked to oxidative stress induced by tissue-specific breakdown of muscle mitochondria [8
Identification of abnormalities in skeletal muscle at the early stage of ALS, preceding the onset of the disease, seems to be important for diagnosis and treatment of patients. Our previous study showed that skeletal muscle in rats bearing the hSOD1 G93A
transgene already showed evidence of oxidative stress at the early stages of disease [9
]. In addition, the ALS rats exhibited an impairment of insulin signaling and iron metabolism in the skeletal muscle that preceded muscle atrophy [11
]. Other symptoms, observed in skeletal muscle, are characteristic for the early stage of disease including the upregulation of proteasome activity and autophagy activation [12
], reduction of mechanical properties of gastrocnemius-soleus muscle group [13
], the impairment of l
channel—a key regulator of both mass and force of skeletal muscle [14
]—loss of forelimb strength (grip test) [15
], abnormal mitochondrial dynamics [16
], and other morphological and functional mitochondria abnormalities [17
Mitochondrial dysfunction and oxidative stress are associated with the development of neurodegenerative diseases [18
]; however, the relationship between these factors and skeletal muscle strength dysfunction at the early stage of ALS is not completely understood.
Mitochondrial dysfunction has been observed in skeletal muscles of hSOD1 G93A
mice, which are related to mutation in SOD1. SOD1 mutations are associated with misfolding and mislocalization of the SOD1 protein. While the normal SOD1 protein is usually found in the cytosol, mutant SOD1 accumulates within mitochondria and appears to contribute to many of the mitochondrial perturbations such as impaired adenosine triphosphate (ATP) production, altered calcium buffering, and initiation of apoptosis [24
]. Luo et al. demonstrated that SOD1 G93A formed aggregates in mitochondria of skeletal muscle, and this aggregation led to mitochondrial depolarization, disruption of the mitochondrial network, and dynamics in skeletal muscle [16
] Moreover, dysfunctional mitochondria can be a source of reactive oxygen species (ROS) generation and cellular oxidative stress [28
]. In addition, Xiao et al. showed an increased ROS generation prior to the onset of ALS symptoms (at the age of 2 months), whereas the onset of ALS symptoms (at the age of four months) was associated with drastic increase in ROS generation [31
]. Decrease in oxidative stress seems to be one of the most important strategies for treating ALS disease and other neuromuscular disorders [32
]. Regulation of metal homeostasis and decrease in oxidative stress, similar to the benefits of swim training, leads to the prolongation of the lifespan of transgenic mice [35
]. Deforges et al. [15
] documented that swim training delays the onset of the first symptoms of the disease, sustains the motor function, and increases the life span of ALS mice by about 25 days. Similar results were obtained in our recent studies [36
]. In contrast to running exercise, large-scale motoneurons are activated by swimming. The swimming-based program is a high frequency exercise associated with high hindlimb movement amplitude (373.9 ± 47.6 cycles min−1
and 4.86 ± 0.40 cm), preferentially activating a sub-population of large motoneurons [37
], which may play an important role in the protection of muscle fibers in ALS. Biochemical and physiological changes induced by exercise have been quite well defined. Data have shown that swimming exercise modifies mitochondria structure and function due to reduction of oxidative stress, improvement of mitochondrial bioenergetics, and regulation of the concentration of mitochondrial cholesterol [36
We hypothesized that at the early stages of the disease, swim training is already a modulator of cellular metabolism via changing muscle bioenergetics, and modulating the oxidative stress can influence the reduction in muscle strength. We also proposed checking which of the parameters are correlated with the reduction in muscle strength in ALS mice, which may be used as potential biomarker of the early phase of the disease. Therefore, the aim of this study was to investigate the effects of swim training and ALS disease on mitochondrial bioenergetics, oxidative stress, and on changes in muscle strength of hSOD1 G93A mice at the early stage of the disease.
The progression of ALS leads to muscle weakness and atrophy, which are associated with deterioration in muscle strength. Therefore, to assess the effects of swim training on the grip strength of ALS mice, we subjected both wild-type (WT) and hSOD1 G93A
mouse groups to a regimen of exercise. Over time, untrained mice lost grip strength as a result of the progression of the disease. According to the expectations, we observed that swim training maintained the grip strength in ALS mice (Figure 1
As we postulated that the decline in the bioenergetics in muscles is a factor favoring the progression of the disease, we decided to measure the activities of individual enzymes or protein complexes involved in oxidative metabolism. The activity of citrate synthase (CS), the first enzyme in the tricarboxylic acid (TCA) cycle, was reduced in the ALS I group of mice (0.385 ± 0.02 μmol/min/mg of protein), compared with the corresponding WT mice (0.540 ± 0.03 μmol/min/mg of protein) (Figure 2
). Additionally, there was a small association of swim training with an increase in CS activity (ALS I SWIM vs. ALS I) (p
= 0.0048, LSD post hoc
test) (Figure 2
The activity of total malate dehydrogenase (MDH) increased in the ALS I and ALS I SWIM groups (7.018 ± 0.21 and 5.982 ± 0.19 μmol/min/mg of protein, respectively) vs. the ALS 0 group (4.300 ± 0.09 μmol/min/mg of protein) (p
= 0.0001) and corresponding WT group of mice (p
= 0.0001). However, swim training was associated with a decrease in MDH activity (ALS I SWIM vs. ALS I) (p
= 0.0003). No changes in MDH enzyme activity were observed in the WT groups of mice (Figure 3
A). The activity of mitochondrial malate dehydrogenase (mMDH) increased in the ALS I and ALS I SWIM groups (4.685 ± 0.20 and 3.904 ± 0.17 μmol/min/mg of protein, respectively) vs. the ALS 0 group (3.265 ± 0.11 μmol/min/mg of protein) and corresponding WT group of mice. However, there was a swim-training-associated decrease in mMDH activity (ALS I SWIM vs. ALS I) (p
= 0.0054). No changes in mitochondrial MDH activity were observed in the WT groups of mice (Figure 3
B). The activity of cMDH increased in the ALS I and ALS I SWIM groups (2.333 ± 0.11 and 2.077 ± 0.11 μmol/min/mg of protein, respectively) vs. the ALS 0 group (1.166 ± 0.08 μmol/min/mg of protein) (p
= 0.0001) and corresponding WT group of mice (p
= 0.0001). The activity of cMDH was higher in WT I vs. WT 0 group of mice (Figure 3
Cytochrome c oxidase (COX) activity increased in the ALS I and ALS I SWIM groups (0.257 ± 0.01 and 0.243 ± 0.01 μmol/min/mg of protein, respectively) vs. the ALS 0 group (0.171 ± 0.01 μmol/min/mg of protein) (p
= 0.0001) and corresponding WT group of mice (p
= 0.0001). No changes in the activity of this enzyme were observed in the WT groups of mice (0.153 ± 0.01, 0.165 ± 0.01, and 0.166 ± 0.01 μmol/min/mg of protein in WT 0, WT I, and WT I SWIM, respectively) (Figure 4
As MDH activity is correlated with COX activity (R
= 0.84, p
= 0.000002), we think that this is a compensatory response, whereby there are no alterations in mitochondrial respiration (Table 1
). The grip strength was correlated only with the activity of CS (R
= 0.68, p
Oxidative stress is another factor that accompanies the development of ALS. Therefore, we decided to measure the concentration of sulfhydryl (SH) groups in the skeletal muscle of ALS and WT mice. In symptomatic ALS mice (ALS I), we observed lower SH groups concentration compared with the ALS 0 group of mice (472.4 ± 23.96 and 579.9 ± 11.57 nmol/g of tissue, respectively) (p
= 0.0019). The level of SH groups was also higher in ALS 0, as compared with the trained ALS mice (497.6 ± 21.92 nmol/g of tissue) (p
= 0.0299) (Figure 5
). There were no significant changes in the level of SH groups in any WT group of mice (586.0 ± 15.30, 552.2 ± 18.54 and 549.0 ± 16.02 nmol/g of tissue in WT 0, WT I, and WT I SWIM, respectively) (Figure 5
). The concentration of SH groups was significantly correlated with COX activity (R
= –0.79, p
4. Materials and Methods
The experiments with animals (project No. 2013/09/NZ7/02538) were approved by the Local Ethics Committee (Resolution No. 11/2013 of 22 April 2013) and in accordance with the approval of the Polish Ministry of the Environment (Decision No. 155/2012).
Transgenic male mice with the G93A human SOD1 mutation B6SJL-Tg (SOD1G93A) 1Gur/J (ALS mice) (n = 24) and wild-type male mice B6SJL (n = 24) were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). The mice were housed in an environmentally controlled room (23 ± 1 °C with a 12 h light/dark cycle); the mice received standard mice chow and water ad libitum. After acclimatization, the mice were randomly divided into the following groups according to disease progression and training status: ALS 0: ALS untrained mice with no visible signs of the disease (n = 8), ALS I: ALS untrained (n = 8), and ALS I SWIM: and ALS trained mice (n = 8). Corresponding groups of wild-type (WT) untrained and trained mice were created: WT 0: WT untrained mice (n = 8), WT I: WT untrained mice (n = 8), and WT I SWIM: WT trained mice (n = 8).
The mice were euthanized by cervical dislocation. The mice from the ALS 0 and WT 0 groups were euthanized on the 70th day of life. The mice from the ALS I group were euthanized when we observed the first symptoms of disease (116 ± 2 days of life). The ALS I SWIM, WT I, and WT I SWIM mice were euthanized at the same age as the ALS I group.
4.2. Swim Training Protocol
Starting at 10 weeks of age, the transgenic (group ALS I SWIM) and the control mice (group WT I SWIM) underwent the training procedure according to Deforges et al. [15
] with slight modification described by Flis et al. [36
4.3. Grip Strength
Grip strength (n = 8 in each group) was tested once a week between 70 and 105 days of life. The time during which the animals were able to sustain their weight holding onto a metal rail suspended midair was recorded, with the maximum time being 180 s. Each mouse was subjected to three trials with a resting period between tests.
4.4. Isolation of Skeletal Muscle Mitochondria
The thigh muscle mitochondria were isolated, as previously described by Makinen and Lee [47
] with slight modifications described by Flis et al. [36
]. The muscles were rapidly removed, trimmed of visible connective tissue, weighed, and placed in 10 mL of ice-cold mitochondrial isolation buffer A (100 mM KCl, 50 mM Tris, 5 mM MgCl2
, 5 mM EDTA, pH 7.4). The muscles were minced with scissors, incubated for 1 min with protease (10 mL of isolation buffer per 1 g of tissue, supplemented with protease (0.2 mg/mL)). After 1 min. of incubation, the same volume of buffer A was added and homogenized using a Teflon pestle homogenizer. The homogenate was centrifuged at 700× g
for 10 min. The supernatant was decanted and centrifuged at 4 000× g
for 10 min. The mitochondrial pellet was resuspended in 30 mL of suspension buffer B (100 mM KCl, 50 mM Tris, 1 mM MgCl2
, 1 mM EDTA, pH 7.4, supplemented with 0.5 % BSA) and centrifuged at 10,000× g
for 10 min. After this washing step had been repeated twice, the final mitochondrial pellet was resuspended in buffer MRB (250 mM mannitol, 5 mM HEPES, 0.5 mM EGTA, pH 7.4) (skeletal muscle mass (mg) × 0.2 mL). All the steps were performed at 4 °C.
4.5. High-Resolution Respirometry
Respiration was measured at 37 °C in a high-resolution respirometer (Oroboros, Oxygraph; Innsbruck, Austria) according to [36
]. Freshly isolated skeletal muscle mitochondria (0.1 mg of mitochondrial protein) were added to 2 mL of the respiration medium (110 mM sucrose, 60 mM K-lactobionate, 0.5 mM EGTA, 0.1 % BSA essentially fatty acid free, 3 mM MgCl2
, 20 mM taurine, 10 mM KH2
, 20 mM K-HEPES, pH 7.1). For the assessment of mitochondrial respiration, the following protocol was used: (1) Non-phosphorylating LEAK respiration was assessed by injecting 2 mM L-malic acid (neutralized with KOH) as NADH (N)-linked substrates, called state NL
; (2) OXPHOS capacity was induced by adding 5 mM ADP and 10 mM of glutamic acid (neutralized with KOH), called state NP
; and (3) OXPHOS coupling efficiency (OCE), calculated with the formula (1 − (state NL
)), reflects the coupling of respiration supported by electron transferring flavoprotein (ETF) with malate as substrate before (state NL
) and after the addition of ADP and glutamic acid (state NP
). The OCE was calculated as a measure of mitochondrial quality and control.
4.6. Measurement of Cytochrome c Oxidase, Citrate Synthase and Malate Dehydrogenase Activities
All enzymes activities were measured spectrophotometrically (Cecil CE9200, Cecil Instruments Limited, Cambridge, UK) in thigh muscles homogenates. The enzymes activities are expressed as μmol/minute/mg of protein. The cytochrome c oxidase (COX) activity was measured at 37 °C, according to Reference [48
]. Briefly, 20 μL of homogenate (1:10, 5%) was incubated for 2 min in 960 μL of buffer (10 mM potassium phosphate buffer, 0.01% Triton-X100, pH 7.2). Next 20 μL of reduced cytochrome c
was added to initiate the reaction. The reactions were conducted in duplicate and absorbance was read at 550 nm.
The citrate synthase (CS) activity was measured at 37 °h in duplicate according to Reference [49
]. Briefly, 10 μL of homogenate (1:10, 5%) was incubated for 2 min in 970 μL of buffer (50 mM Tris-HCl, 1 mM EDTA, 0.01% Triton-X100, pH 7.8) supplemented with 10 μL of freshly made DTNB (10 mM) and 10 μL acetylCoA (50 mM). Next 10 μL of freshly made oxaloacetic acid (10 mM) was added to initiate the reaction. The reactions were conducted in duplicate and absorbance was read at 412 nm.
The malate dehydrogenase (MDH) activity was measured at 30 °C according to [50
]. Briefly, 10 μL of homogenate (1:10, 5%) was incubated for 2 min in 970 μL of buffer (50 mM Tris-HCl, 5 mM EDTA, 0.01% Triton-X100, pH 7.6) supplemented with 10 μL of freshly made NADH (20 mM). Next 8.5 μL of freshly made oxaloacetic acid (20 mM) was added to initiate the reaction. The reactions were conducted in duplicate and absorbance was read at 340 nm. The mitochondrial MDH (mMDH) activity was assessed after 2 min preincubation of 20 μL homogenate (1:10, 5%) with 98% ethanol (1:1 vol). The cytosolic MDH (cMDH) was calculated as the difference between MDH and mMDH activities.
4.7. Manifestation of Oxidative Stress
The sulfhydryl groups (SH) [51
] were measured in the skeletal muscles homogenates. Briefly, 20 μL of homogenate (5%) was added to 200 μL of buffer (10 mM potassium phosphate buffer, pH 8.0). Next 30 μL of SDS (10 %) and 30 μL of DTNB (1 mM) were added. The measurements were conducted in duplicate. After this, the probes were mixed and incubated for 30 min in 37 °C. The absorbance was measured at 412 nm in microplate reader Thermo Scientific Multiscan Go (ThermoFisher Scientific, Vartaa, Finland). The values of the sulfhydryl groups (mmol/g of tissue) were calculated against sample blank (without DTNB) from the standard curve of reduced glutathione.
4.8. Data Analysis
Statistical analyses were performed using a software package Statistica v. 13.0 (StatSoft Inc., Tulsa, OK, USA). The results are expressed as the mean ± standard error (SE). The differences between the group means for tissue-based assays were performed using two-way ANOVA. The changes in mean scores of grip strength over time points were tested using the repeated measures ANOVA. If a difference was detected in the ANOVA model, the significant differences were determined using the Tukey’s post-hoc test. To verify the significance of the small, swim training associated changes (ALS TER vs. ALS SWIM), the Least Significant Difference (LSD) post-hoc test was used. The results were considered statistically significant when p < 0.05. A Pearson product-moment correlation coefficient was computed to assess the relationship between obtained results.