Subcellular distribution of glycogen and decreased tetanic calcium in fatigued single intact mouse muscle fibres

Since the turn of the century the physiological and metabolic regulators of fatigue within skeletal muscle have been extensively researched (Hill, 1925). It is well established that calcium (Ca²⁺), released from the sarcoplasmic reticulum (SR), plays an integral role in skeletal muscle contractile function through the initiation of cross bridge cycling. It has been observed in vitro that SR Ca²⁺ release reduces during the onset of fatigue, which is believed to explain the subsequent loss of force generation within skeletal muscle. A series of recent ex vivo experiments have shown that the loss of Ca²⁺ release from the SR is paralleled with a reduced availability of endogenous skeletal muscle glycogen stores (Ørtenblad et al. 2013). Following this observation, there have been suggestions that the onset of fatigue within isolated skeletal muscle is controlled by the subcellular localisation of glycogen, which would indicate a relationship to SR Ca²⁺ release. However to date, no study has used an integrated model to investigate whether the locality of skeletal muscle glycogen influences fatigue and the relationship this has to SR Ca²⁺ release.

In a recent edition of The Journal of Physiology, Nielsen et al. (2014) performed a novel and elegant integrated series of studies to investigate the role of glycogen depletion on Ca²⁺ release and power output within an isolated single mouse muscle fibre. The fibres were divided into one of four groups: control, high intensity fatigue (HIF), low intensity fatigue (LIF) and low intensity equivalent (LIE). All groups underwent the same electrical stimulation using 70 Hz until fatigue, categorised as a natural force reduction of 70%. The interval (rest) between each stimulation varied according to their treatment. For HIF, 70 Hz was repeated every 2 s, while for LIF 70 Hz was repeated every 10 s, until fatigue. LIE was stimulated every 10 s for 42 contractions (the mean number of stimulations for the HIF group). This approach is advantageous as it provides a model looking at varying exercise intensities that somewhat mimics maximal and sub-maximal exercise. Throughout the stimulation, Ca²⁺ release was measured using Indo-1 fluorescence signals via a photomultiplier tube. Following the onset of fatigue, glycogen content was measured using electron microscopy. Subcellular fractions were quantified using transmission electron microscopy and stereological methods. The volume fraction was calculated by taking into account each subceullular section area (estimated by point counting), thickness and diameter.

The authors report that during fatiguing stimulation Ca²⁺ release significantly fell, with the fall being steeper for HIF compared to LIF which was paralleled with a similar decrement of force production. Overall the functional changes to acute fatigue in the fibres appeared larger for the HIF with a slowing of relaxation, a more preserved Ca²⁺ release at the point of fatigue and a larger rise in resting [Ca²⁺]_i after fatigue when compared to LIF. On the other hand, the LIE fibres also appeared to lose force during the tetani but were divided into two groups that were either fatigued (LIE-F > 75% force loss) or non-fatigued (LIE-NF < 50% force loss). As expected the LIE-F observed a large reduction in force and calcium release whereas LIE-NF did not. Alongside the functional differences in the muscle fibres, glycogen content fell at fatigue for all fibres in each group with similar reductions observed for intramyofibrillar and intermyofibrillar glycogen pools in HIF and LIF. Subsarcolemmal glycogen content was not lower in any group of fibres when compared to control. Taking the Ca²⁺ release and glycogen content data together, a strong relationship appeared to exist between decreased tetanic [Ca²⁺]_i and low glycogen with a closer association being observed for intra- (R² = 0.37, P = 0.0008) compared to inter- (R² = 0.16, P = 0.051) myofibrillar glycogen depletion. In contrast, no relationship existed between subsarcolemmal glycogen depletion and Ca²⁺ release. Collectively, these new data propose that subcellular glycogen content, specifically of the intramyofibrillar pool, modulates Ca²⁺ release from the SR and ultimately fatigue.

This study implemented a well-planned and controlled model to demonstrate that subcellular glycogen depletion influences SR Ca²⁺ release, at least in vitro. The novel data on compartmental glycogen depletion also provide support for the intriguing hypothesis of metabolic microenvironments possibly dictated by glycogen within cells. The authors should be commended for their work having taken the first logical step in a series of experiments needed to understand the relationship between glycogen depletion and fatigue. The intriguing data presented by Nielsen et al. (2014) do, however, raise a number of new questions.

The in vitro methodology presented by the authors uses a glucose-free solution to minimise metabolic variability; however, this does obscure certain metabolic mechanisms that might be present in vivo from being elucidated. Therefore further research should repeat this model using an experimental medium with physiologically relevant metabolite availability so that the influence of free glucose can also be taken into consideration. Also, it is unlikely that the model fully isolated the role of glycogen depletion on SR Ca²⁺ release as functional differences between stimulation protocols were apparent. After stimulation, HIF fibres had a greater repletion of calcium, a slowing of relaxation and, at fatigue, a more preserved calcium release suggesting force decreases were partly due to decreased calcium sensitivity. Similarly, it is not yet clear whether glycogen depletion from differing compartments has differing effects on fatigue. This paper found a correlation between intra- and intermyofibrillar glycogen depletion and SR Ca²⁺ release, but not subsarcolemmal; however, based on the current study, at control these compartments store 72%, 28% and 4% of glycogen, respectively. Thus a lack of significant correlation in subsarcolemmal glycogen depletion may reflect a lower statistical power and not necessarily a reduced influence on SR Ca²⁺ release. It may be that glycogen depletion, regardless of compartment, influences SR Ca²⁺ release. A final question raised by this work is whether a threshold value exists before glycogen depletion influences SR Ca²⁺ release and whether this translates to different mammalian exercise models. In this study, glycogen depletion >80% of control was required for a consistent negative impact on SR Ca²⁺ release but this level of depletion is extremely uncommon for exercising humans. Furthermore, this model only used type II mouse fibres which are highly glycolytic, whereas type I fibres support exercising tissue primarily through oxidative metabolism of fats. Fatigue in these fibres is probably regulated by multiple mechanisms rather than glycogen depletion alone, such as Na⁺–K⁺-ATPase activity, so this relationship may be less relevant in a whole muscle with varied fibre composition and substrate utilisation. Together these questions mask the extent to which glycogen depletion influences sarcoplasmic Ca²⁺ release in whole exercising muscle in mice, or humans especially as mouse type II fibres have only 20% of the glycogen content of type II human muscle fibres.

With these questions in mind, it would be fascinating to extend this research into human exercise models. With advancing techniques, single fibre approaches are now becoming increasingly common in human experimentation (Murphy, 2011) and so repeating this model on isolated human type II muscle fibres would provide the first logical translation of this work. This could be followed by repeating for human type I fibres and then for whole muscle, providing a comprehensive analysis of the relationship between glycogen depletion and fatigue, in each muscle fibre type and whole muscle, in humans. Further examination could be supported using methodologies to quantify regional glycogen depletion, such as the immunofluorescence approach recently demonstrated by Prats et al. (2013). This approach not only allows the simultaneous examination of glycogen and intramuscular triacylglycerol stores, but also provides more accuracy than the PAS stain used by Nielsen et al. (2014), which has the drawback of also binding to glycoproteins and proteoglycans and so is not truly glycogen specific. Finally, nutrition/exercise-based interventions could then also be used to manipulate regional glycogen stores and examine skeletal muscle fatigue.

In conclusion, the study by Nielsen et al. (2014) provides important novel insights into the role of compartmental glycogen depletion on calcium release and fatigue in skeletal muscle. These data are integral to advancing the field of exercise physiology and metabolism in order to understand the functional implications of glycogen for human health, but also exercise performance to include regulating substrate metabolism, metabolic flexibility and skeletal muscle function. Whilst fatigue is undoubtedly a complex multi-factorial process, a deeper understanding of the mechanisms and impacts of glycogen depletion in each compartment will help complete understanding and could eventually support more specific exercise and nutrition interventions to modulate skeletal muscle metabolism and fatigue. This is highly relevant to athletes, but through further work could also be relevant to those with impaired metabolic health, including the elderly, obese and chronic disease models such as diabetes or cardiovascular disease. The authors should once again be praised for presenting novel data with an elegant methodology that has progressed our understanding of skeletal muscle metabolism.

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Competing interests

The authors confirm that they have no conflicts of interest to disclose.