Altered mitochondrial bioenergetics and ultrastructure in the skeletal muscle of young adults with type 1 diabetes
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A comprehensive assessment of skeletal muscle ultrastructure and mitochondrial bioenergetics has not been undertaken in individuals with type 1 diabetes. This study aimed to systematically assess skeletal muscle mitochondrial phenotype in young adults with type 1 diabetes.
Physically active, young adults (men and women) with type 1 diabetes (HbA1c 63.0 ± 16.0 mmol/mol [7.9% ± 1.5%]) and without type 1 diabetes (control), matched for sex, age, BMI and level of physical activity, were recruited (n = 12/group) to undergo vastus lateralis muscle microbiopsies. Mitochondrial respiration (high-resolution respirometry), site-specific mitochondrial H2O2 emission and Ca2+ retention capacity (CRC) (spectrofluorometry) were assessed using permeabilised myofibre bundles. Electron microscopy and tomography were used to quantify mitochondrial content and investigate muscle ultrastructure. Skeletal muscle microvasculature was assessed by immunofluorescence.
Mitochondrial oxidative capacity was significantly lower in participants with type 1 diabetes vs the control group, specifically at Complex II of the electron transport chain, without differences in mitochondrial content between groups. Muscles of those with type 1 diabetes also exhibited increased mitochondrial H2O2 emission at Complex III and decreased CRC relative to control individuals. Electron tomography revealed an increase in the size and number of autophagic remnants in the muscles of participants with type 1 diabetes. Despite this, levels of the autophagic regulatory protein, phosphorylated AMP-activated protein kinase (p-AMPKαThr172), and its downstream targets, phosphorylated Unc-51 like autophagy activating kinase 1 (p-ULK1Ser555) and p62, was similar between groups. In addition, no differences in muscle capillary density or platelet aggregation were observed between the groups.
Alterations in mitochondrial ultrastructure and bioenergetics are evident within the skeletal muscle of active young adults with type 1 diabetes. It is yet to be elucidated whether more rigorous exercise may help to prevent skeletal muscle metabolic deficiencies in both active and inactive individuals with type 1 diabetes.
KeywordsCalcium tolerance Capillary density Mitochondria Myopathy Reactive oxygen species Skeletal muscle Type 1 diabetes
AMP-activated protein kinase
Ca2+ retention capacity
Mitochondrial glycerol-3-phosphate dehydrogenase
Mitochondrial permeability transition pore
Magnetic resonance spectroscopy
Oxidative phosphorylation cocktail
Pyruvate dehydrogenase complex
Reactive oxygen species
Transmission electron microscopy
Unc-51 like autophagy activating kinase 1
Skeletal muscle accounts for almost half of our body weight and, as such, is responsible for up to 75% of insulin-stimulated glucose disposal after a meal [1, 2]. In fact, blood glucose uptake into muscle and its subsequent conversion to glycogen is a major determinant of insulin sensitivity [3, 4]. Given the importance of skeletal muscle for glucose management, as well as its vital role in insulin action, impairments in skeletal muscle health in people with type 1 diabetes diminishes their ability to mitigate dysglycaemic burdens, promotes the development and progression of complications [5, 6] and, ultimately, expedites the accelerated physical disability that characterises diabetes . However, our understanding of the impact of this chronic disease on the health/quality of human skeletal muscle is extremely limited.
Mitochondria play a fundamental role in energy metabolism and ATP production throughout the body. Based on their role in fuel oxidation, mitochondria can also generate considerable amounts of reactive oxygen species (ROS). It remains controversial whether impaired skeletal muscle mitochondrial respiratory function contributes to insulin resistance and dysglycaemia in obesity and type 2 diabetes . However, a consistent observation has been increased mitochondrial ROS in obese/type 2 diabetes relative to lean control individuals . In addition, using 31P-magnetic resonance spectroscopy (MRS), people with type 1 diabetes have been shown to exhibit slowed post-exercise ATP resynthesis [9, 10, 11]; the underlying mechanism(s) for this impairment is currently unknown and would require a direct and comprehensive exploration of site-specific mitochondrial respiratory function. Moreover, the effect of type 1 diabetes on skeletal muscle ultrastructure, last reported in 1977 , has not been re-examined since the introduction of more aggressive insulin therapies (e.g. basal and fast-acting insulins, insulin analogues and pump therapy), and its effect on muscle mitochondrial ROS production remains to be elucidated.
Here, we hypothesised that otherwise healthy, young adults with type 1 diabetes would display site-specific decrements in skeletal muscle mitochondrial respiration, increased ROS production and altered muscle ultrastructure relative to non-diabetic controls.
Demographics of participants
Control (n = 12)
Type 1 diabetes (n = 12)
26 ± 2
26 ± 4
65.4 ± 12.1
73.0 ± 13.4
1.70 ± 0.07
1.69 ± 0.13
22.5 ± 2.8
25.4 ± 3.7*
33.0 ± 2.2
63.0 ± 16.0***
5.2 ± 0.2
7.9 ± 1.5***
Diabetes duration (years)
15.2 ± 7.9
Diabetes onset (years of age)
11.5 ± 7.3
Physical activity (min/week)a
233.1 ± 152.6
201.0 ± 168.6
Participants reported to York University in the early morning or early afternoon and were instructed to consume a standardised meal 1.5–2 h prior to their visit. Participants with type 1 diabetes were also instructed to continue their habitual use of insulin. Upon arrival, body mass and height measurements were taken to determine BMI, and a blood sample was obtained using venepuncture, for HbA1c analysis. Skeletal muscle samples were then obtained from the vastus lateralis muscle by a microbiopsy percutaneous needle, as described previously , and used for mitochondrial bioenergetic analyses, transmission electron microscopy (TEM), histological analysis and western blotting, as described below.
All mitochondrial bioenergetic experiments were performed in vitro using permeabilised muscle fibres prepared from fresh muscle samples that were immediately placed in ice-cold BIOPS containing (in mmol/l): 50 MES, 7.23 K2EGTA, 2.77 CaK2EGTA, 20 imidazole, 0.5 dithiothreitol (DTT), 20 taurine, 5.77 ATP, 15 PCr and 6.56 MgCl2.6H2O (pH 7.1), as previously described in detail .
Using the Oxygraph-2k (Oroboros Instruments, Innsbruck, Austria), high-resolution measurements of mitochondrial oxygen consumption were conducted in permeabilised myofibres placed in 2 ml of respiration medium MiR05 , containing 0.5 mmol/l EGTA, 10 mmol/l KH2PO4, 3 mmol/l MgCl2.6H2O, 60 mmol/l K-lactobionate, 20 mmol/l Hepes, 20 mmol/l taurine, 110 mmol/l sucrose and 1 mg/ml fatty acid-free BSA (pH 7.1), at 37°C. Experiments were conducted at an initial oxygen concentration of 350–375 μmol/l with constant stirring at 750 rev/min. Respiration medium was supplemented with 20 mmol/l creatine (C0780; Sigma, St Louis, MO, USA) to enhance mitochondrial phosphate shuttling . ADP-stimulated respiratory kinetics were determined through ADP (A5285; Sigma) titrations in the presence of 5 mmol/l blebbistatin (CAY13013; Cayman Chemical, Ann Arbor, MI, USA), in order to prevent spontaneous contraction of fibres [13, 17, 18], 5 mmol/l pyruvate (P2256; Sigma) and 2 mmol/l malate (M1000; Sigma). Complex I kinetics were determined through standard pyruvate and glutamate (GLU303; BioShop, Burlington, ON, Canada) titrations in the presence of 5 mmol/l blebbistatin, 5 mmol/l ADP and 2 mmol/l malate, while Complex II kinetics were determined through standard succinate (S2378; Sigma) titrations in the presence of 5 mmol/l blebbistatin, 5 mmol/l ADP and 10 μmol/l rotenone (R8875; Sigma; to prevent superoxide generation at Complex I). During each titration, Cytochrome c (192-10; Lee Biosolutions, Maryland Heights, MO, USA) was added last to determine intactness of the outer mitochondrial membrane. Any samples that exceeded a 10% increase in respiration after Cytochrome c addition were excluded .
Mitochondrial H2O2 emission
Mitochondrial H2O2 emission was measured fluorometrically (QuantaMaster 40; HORIBA Scientific, Edison, NJ, USA) at 37°C in buffer Z , containing 105 mmol/l K-MES, 30 mmol/l KCl, 10 mmol/l KH2PO4, 5 mmol/l MgCl2.6 H2O, 1 mmol/l EGTA, 5 mg/ml BSA (pH 7.1), supplemented with 20 mmol/l creatine, 10 μmol/l Amplex UltraRed reagent (A36006; Life Technologies, Carlsbad, CA, USA), 0.5 U/ml horseradish peroxidase (P8375; Sigma) and 40 U/ml Cu/Zn superoxide dismutase (SOD1; S9697; Sigma), as described previously . Various sites were assessed by titrating the following substrates [20, 21]: (1) 2.5 μmol/l antimycin (A8674; Sigma) for assessment of Complex III; (2) 10 mmol/l pyruvate and 2 mmol/l malate, to assess Complex I via generation of NADH; (3) 10 mmol/l succinate, to assess Complex I (via reverse flow) via generation of FADH2; (4) 10 mmol/l pyruvate and 0.5 μmol/l rotenone, to assess pyruvate dehydrogenase complex (PDC); rotenone blocks NADH entry into Complex I; and (5) 20 mmol/l glycerol-3-phosphate, for the assessment of mitochondrial glycerol-3-phosphate dehydrogenase (mG3PDH; G7886; Sigma). In addition, fibres used for Complex I (pyruvate + malate), PDC and mG3PDH measurements were treated with 35 μmol/l 1-chloro-2,4-dinitrobenzene (237329; Sigma) during permeabilisation to deplete glutathione .
Mitochondrial Ca2+ retention capacity
Mitochondrial Ca2+ retention capacity (CRC) was measured in duplicate, fluorometrically (QuantaMaster 40; HORIBA Scientific), at 37°C in buffer Y, containing 250 mmol/l sucrose, 10 mmol/l Tris-HCl, 20 mmol/l Tris base, 10 mmol/l KH2PO4, 2 mmol/l MgCl2.6H2O and 0.5 mg/ml BSA, with 1 μmol/l Calcium Green-5N (C3737; Life Technologies), as described previously . Briefly, Ca2+ uptake was initiated with 8 nmol pulses of Ca2+ (CaCl2) and then subsequent pulses of 4 nmol Ca2+ were titrated until opening of the mitochondrial permeability transition pore (mPTP) was observed .
Fresh muscle was immediately fixed in 2% (vol./vol.) glutaraldehyde (111-30-8; Electron Microscopy Sciences, Hatfield, PA, USA) in 0.1 mol/l sodium cacodylate buffer pH 7.4 (Canemco, Lakefield, QC, Canada) and processed as described previously . To quantify mitochondria, representative micrographs from eight unique fibres (containing a portion of the subsarcolemmal region adjacent to the nucleus, with most of the image containing the intermyofibrillar area) were acquired at ×15,000 magnification. Blinded quantification of mitochondrial size (mean area, μm2), distribution (number per μm2) and density (μm2 × number per μm2 × 100) was achieved using Nikon Imaging Software (NIS)-Elements AR (v 4.6; Nikon, Melville, NY, USA) by manually outlining mitochondria and converting to actual size using a calibration grid .
Samples were oriented and trimmed to allow longitudinal sectioning of fibres, which were visually localised. Ultra-microtomy (Leica UC7; Leica Microsystems, Wetzlar, Germany) was performed with a 35° diamond knife (Diatome, Nidau, Switzerland), and 65 nm thick sections were mounted onto copper slot grids (Plano, Wetzlar, Germany). Electron micrographs for tomography tilt series were acquired on a JEM-2200FS electron microscope (JEOL, Akishima, Tokyo, Japan) (200 kV) equipped with a TemCam-F416 camera (TVIPS, Munich, Germany) and using a 25 electron volt (eV) energy filter slit. Under-focus was adjusted to 4000 nm. Micrographs were acquired two-fold binned for tomography. The resulting pixel size was 2.4 nm. Tilt series was collected over a total angular tilt range from −70° to +70°, at 2° increments. Image series was aligned by patch tracking mode using the IMOD software package (v 4.7; http://bio3d.colorado.edu/imod) . A single reconstructed volume was computed from each tilt series by radially weighted back projection. A 3D density distribution (tomogram) was obtained. A virtual section with a 24 nm thickness was generated with the slicer tool in IMOD. Major organelles were coloured utilising Adobe Photoshop CC 2015 by a blinded researcher.
Histology and immunofluorescence analysis
Biopsied muscle was immediately embedded in Tissue Tek optimum cutting temperature (O.C.T.) compound (4583; VWR, Radnor, PA, USA), frozen in liquid nitrogen-cooled isopentane and stored at −80°C until analysis. Cryosections (−20°C) that were 8 μm thick were transferred onto static-free microscope slides (CA48311-703; VWR), allowed to air dry and fixed with ice-cold 4% (wt/vol.) paraformaldehyde. Sections were then incubated in blocking solution (5% [vol./vol.] normal goat serum in 1X PBS; S-1000; Vector Laboratories, Burlingame, CA, USA) for 30 min and then co-stained with CD31 (platelet endothelial cell adhesion molecule-1 [PECAM-1]) overnight. An antibody against CD41 was added the following day and incubated for 1 h. The appropriate secondary antibodies were then applied for 1 h: Alexa Fluor 594 and Alexa Fluor 488. Nuclei were counterstained with DAPI. CD31 was used as a marker of capillary content and CD41 to measure platelet aggregation. The antibodies were validated by manufacturers for immunofluorescence analyses (see electronic supplementary material [ESM] Table 1 for antibody details). H&E stains were used for the determination of muscle fibre area in order to calculate capillary:fibre ratio, as previously described . Images were captured with a Nikon 90 Eclipse microscope (Nikon) and analysed using NIS-Elements AR software (v 4.6; Nikon) at original magnification of ×40.
A portion of muscle was quickly snap-frozen and stored at −80°C until analysis. Frozen muscle (~10–30 mg) was homogenised with a tapered teflon pestle in ice-cold buffer containing 40 mmol/l Hepes, 120 mmol/l NaCl, 1 mmol/l EDTA, 10 mmol/l NAHP2O7·10H2O pyrophosphate, 10 mmol/l β-glycerophosphate, 10 mmol/l NaF and 0.3% CHAPS detergent (pH 7.1, adjusted using KOH). Protein concentrations were determined using a BCA assay (Life Technologies) and then loaded equally (50 or 100 μg), using Ponceau S (P7170; Sigma) as a loading control. Proteins were separated by SDS-PAGE, transferred to polyvinylidene fluoride membrane, blocked for 1 h and incubated overnight at 4°C using commercially available antibodies validated by manufacturers for western blotting: human total oxidative phosphorylation cocktail (OXPHOS; to detect individual complexes of the electron transport chain [I, II, III, IV, V]), phosphorylated (Thr172) and total AMP-activated protein kinase (AMPKα), phosphorylated (Ser555) and total Unc-51-like autophagy activating kinase 1 (ULK1), sequestosome (SQSTM1; herein referred to as p62) and vinculin. After overnight incubation, membranes were washed in tris-buffered saline, 0.1% (vol./vol.) Tween 20 (TBS-T) and incubated for 1 h at room temperature with anti-mouse infrared fluorescent secondary antibody (IRDye 800CW-conjugated) for OXPHOS, and horseradish peroxidase conjugated secondary antibody for all other proteins (see ESM Table 1 for antibody details). Immunoreactive proteins were detected by infrared imaging (Odyssey CLx; LI-COR Biosciences, Lincoln, NE, USA) or chemiluminescence (Gel Logic 6000 Pro Imager; Carestream, Rochester, NY, USA) and quantified by densitometry using Image J (v 1.51t; http://imagej.nih.gov/ij/).
All results are expressed as mean ± SEM except for participant characteristics, which are expressed as mean ± SD. An unpaired two-tailed Student’s t test was performed for all experiments except for CRC and mitochondrial respiration kinetics where an unpaired one-tailed t test and a two-way ANOVA were used, respectively. Significance was established at p ≤ 0.05.
Skeletal muscle mitochondrial bioenergetics
Control of oxidative phosphorylation by ADP
We then determined if mitochondrial sensitivity to specific substrates was different in the muscle of those with type 1 diabetes vs control individuals. No detectable change in Complex I-supported mitochondrial respiration was observed, as evidenced by the lack of a significant difference in the sensitivity or capacity of the mitochondria to oxidise pyruvate (an index of glucose oxidation) (Fig. 1b). This suggests that mitochondrial oxidation of pyruvate from glucose is not impaired in type 1 diabetes, and neither is Complex I oxidation of NADH from pyruvate dehydrogenase. There were also no detectable changes in glutamate-supported respiration (NADH oxidation) (Fig. 1c), which further suggests there is no deficiency in Complex I activity in type 1 diabetes. In contrast, Complex II-supported respiration by succinate (FADH2 synthesis) was significantly lower (p < 0.001, main effect) at both low (sensitivity) and maximal (oxidative capacity) succinate concentrations in participants with type 1 diabetes, suggesting a reduction in Complex II activity in type 1 diabetes compared with control individuals (Fig. 1d).
Mitochondrial H2O2 emission
Next, we measured mitochondrial H2O2 emission potential. In the participants with type 1 diabetes, Complex III-supported mitochondrial H2O2 emission was significantly elevated (p < 0.05) compared with control participants but was not statistically different at Complex I (reverse [succinate] and forward electron flow [pyruvate + malate]), PDC and mG3PDH (Fig. 1e).
Skeletal muscle mitochondrial content
Skeletal muscle mitochondrial Ca2+ tolerance and electron tomography
We next tested Ca2+ handling within the mitochondria. Opening of the mPTP has been linked to swelling and energetic dysfunction . We hypothesised that reductions in Ca2+ handling within the mitochondria (i.e. reduced CRC) would lead to an increased sensitisation of the mPTP and, ultimately, attenuations in mitochondrial respiratory function. As hypothesised, mitochondrial CRC was significantly lower (p = 0.046) in skeletal muscle from participants with type 1 diabetes relative to control participants (Fig. 3a).
We next used electron tomography to assess the skeletal muscle ultrastructure. This powerful technique allows for the 3D study of cellular architecture at the nanoscale level. Consistent with our observed attenuations in mitochondrial respiration, electron tomography revealed irregularities in the organisation of cristae in the participants with type 1 diabetes that were uncommon in control participants (Fig. 3b; mitochondria and cristae highlighted in red). Furthermore, consistent with the reductions in mitochondrial Ca2+ handling previously noted, the muscles from participants with type 1 diabetes also exhibited an increase in size and number of autophagic remnants compared with the control group, and this was visible within both the subsarcolemmal and intermyofibrillar areas (Fig. 3c; highlighted in purple and cyan, respectively). A close association of autophagic remnants to intramyocellular lipid droplets was also frequently observed in individuals with type 1 diabetes (Fig. 3c; lipid droplets highlighted in green). Despite these observations, no differences in the autophagic regulatory protein p-AMPKαThr172 (as noted previously) or its downstream targets p-ULK1Ser555 or p62 were observed between groups (Fig. 3d–f).
Skeletal muscle microvasculature
The findings of this work highlight mitochondrial and autophagic differences within the muscles of young adults with type 1 diabetes with moderate glycaemic control and who exceed recommended activity levels (>150 min of moderate-to-vigorous intensity activity per week) according to the American and Canadian diabetes associations [30, 31]. Specifically, the work described herein demonstrates, for the first time, site-specific alterations within the mitochondria of young adults with type 1 diabetes compared with (matched) control participants without diabetes. These alterations in mitochondrial bioenergetics occur in the absence of a loss of mitochondrial content or changes in skeletal muscle microvasculature (as assessed by capillary density and platelet aggregation). Furthermore, TEM investigations revealed that type 1 diabetes negatively affects skeletal muscle ultrastructure, as evidenced by disorganised mitochondrial cristae and an increased presence of autophagic remnants. These early alterations precede the well-characterised complications (e.g. cardiovascular disease, polyneuropathy) of type 1 diabetes and occur in large, proximal muscle groups (in contrast to distal limb muscles, which are more readily affected by polyneuropathy), suggesting that the early alterations seen in this study are unlikely to be owing to neuropathy but, instead, intrinsic alterations in muscle, and that this skeletal muscle metabolic dysregulation is a primary outcome of this chronic disease that may contribute to dysglycaemia.
Skeletal muscle is not only the primary organ system responsible for physical capacity but is also a critical metabolic organ since it is: (1) the major site of fatty acid catabolism ; (2) a principal mediator of whole-body glucose homeostasis ; and (3) a major determinant of whole-body insulin sensitivity [2, 3]. Thus, impairments in skeletal muscle quality, particularly in conditions of metabolic disease (e.g. diabetes mellitus), could have profound long-term consequences by increasing the incidence of primary contributors to the development of diabetic complications, such as recurrent dysglycaemia, dyslipidaemia and insulin resistance [33, 34, 35]. Consistent with the importance of skeletal muscle in overall health, if one considers other chronic diseases (e.g. heart disease and cancer), loss of muscle mass, strength and metabolic function (referred to as cachexia) are commonly observed, and this decline in muscle health is an important determinant in overall survival [36, 37].
While changes in skeletal muscle health of rodent models of type 1 diabetes has been more intensely studied (previously referred to as ‘diabetic myopathy’ [38, 39]), the extent to which this is happening in human skeletal muscle of those with type 1 diabetes has yet to be fully elucidated . In the clinical diabetes literature, skeletal muscle mitochondrial ATP synthesis is one of the most widely investigated variables, often assessed indirectly in vivo by monitoring the resonance of phosphate in ATP using 31P-MRS. Previous studies using 31P-MRS have frequently, but not always , reported a ~20–25% reduction in mitochondrial ATP synthesis rates in the skeletal muscle of those with type 1 diabetes [9, 10, 11]. While 31P-MRS has the obvious advantage of capturing ATP kinetics under intact physiological conditions, a drawback of this approach is the lack of mechanistic understanding underlying the changes in mitochondrial function that may be observed.
Consistent with the work of Cree-Green et al  and others [10, 11], we observed a ~20% reduction in mitochondrial oxidative capacity in the muscles of those with type 1 diabetes (Fig. 1a, Max), suggesting that our in vitro (permeabilised muscle fibre method) assessment is consistent with 31P-MRS studies. What was unanswered from the aforementioned 31P-MRS studies, however, was whether the mitochondria are reduced in content and/or impaired, or whether nutrient delivery is the cause of the resulting poor mitochondrial function in those with type 1 diabetes. Here we provide the mechanistic insight to answer these questions.
First, regarding whether the deficits observed in previous 31P-MRS studies were directly related to differences in mitochondrial content or impairments of mitochondrial quality, we now offer invaluable insight using ultrastructural analysis. Using TEM, we observed no difference in the amount, distribution and size of mitochondria within the skeletal muscle of those with type 1 diabetes compared with control individuals. Within the mitochondria, we observed no differences in the protein expression of the individual electron transport chain protein subunits using western blot analysis (total OXPHOS cocktail), further suggesting that mitochondrial content is not reduced in young, physically active adults with type 1 diabetes vs control individuals. In contrast, we did observe structural abnormalities in mitochondria using electron tomography; disorganised cristae and more abundant autophagic structures were present in the muscles of individuals with type 1 diabetes compared with control participants. In line with this, we also observed a significant reduction in mitochondria viability in permeabilised myofibres of those with type 1 diabetes, as assessed by CRC .
Second, these findings suggest that nutrient delivery in the muscles of young adults with type 1 diabetes is not likely impaired. Histological/immunofluorescent examination of skeletal muscle biopsies revealed that capillary density within the vastus lateralis muscle was not different between those with type 1 diabetes and those without. Furthermore, as increases in prothrombotic factors have been reported in those with type 1 diabetes , we investigated whether the capillaries present were occluded by platelet aggregates (thrombi) and found that they were not. While we do not see visible evidence for impaired blood flow in the present study, we cannot rule out the possibility of a functional impairment resulting in reduced blood flow and delivery of oxygen and nutrients within the muscles of those with type 1 diabetes .
Collectively, these novel results provide evidence of direct alterations in skeletal muscle mitochondria in those with type 1 diabetes. The underlying mechanism(s) for these mitochondrial alterations in type 1 diabetes may be revealed by the mitochondrial bioenergetics investigations related to mitochondrial H2O2 emission in this study. To the best of our knowledge, mitochondrial-derived ROS production has never been examined in skeletal muscle of humans with type 1 diabetes. We observed a significant increase in mitochondrial H2O2 emission potential at the level of Complex III, with no significant differences owing to Complex I, intermembrane (mG3PDH) or matrix (PDC) proteins. Work using isolated mitochondria [43, 44], has demonstrated that Complex III significantly contributes to ROS production in response to fatty acid oxidation. While we did not assess this in the current study, future work should be undertaken to discern differences in the propensity for ROS production at Complex III in response to glucose and lipid substrate availability in those with type 1 diabetes. The tissue requirements for the comprehensive assessment of mitochondrial phenotype prohibited additional assessment of major mitochondrial antioxidant enzymes (e.g. glutathione peroxidase, thioredoxin reductase and superoxide dismutase) and markers of oxidative stress in the present study. Nonetheless, it has been suggested that Complex III-generated ROS serves as a signalling molecule, particularly in the adaptive response to hypoxia (for review see ). This latter point may be physiologically important when considering the peripheral vascular disease that is known to develop in older adults with type 1 diabetes.
This investigation revealed that young, active people with type 1 diabetes possess mitochondrial and muscle ultrastructural abnormalities that precede reductions in capillary content. While we have demonstrated site-specific alterations in mitochondrial bioenergetics, further investigation is required to identify the precise signalling pathway leading to these changes. Though our observations of increased autophagic remnants in muscles of those with type 1 diabetes were not associated with increases in autophagic markers (p-ULK1Ser555 and p62), it is possible that these signals had already been transmitted and pathways initiated. Alternatively, recent work  has suggested that p62 activity is redox sensitive and may promote autophagy in the absence of changes in content. Clearly, future work in this area will help further elucidate this important observation.
It seems likely that the relationship between skeletal muscle abnormalities and dysglycaemia in type 1 diabetes are reciprocal, particularly given previous suggestions that excessive glucose provision to mitochondria elevates ROS production [47, 48]. Another possible mechanism may relate to peripheral insulin administration. Specifically, in people without type 1 diabetes, large rises in blood glucose levels following a glucose load are attenuated by delivering insulin to the liver first via the portal circulation. However, in those with type 1 diabetes, peripheral insulin administration bypasses the canonical ‘liver-first’ model and results in greater elevations in circulating blood glucose . The peripheral insulin administration shifts a greater burden of the glycaemic load to skeletal muscle, forcing it to accept a greater glycaemic load then it otherwise would [1, 2], thereby increasing glycaemic stress akin to that previously described  for cell types that could not regulate cellular glucose entry. An important avenue for future research is to investigate this altered relationship between blood glucose levels, peripheral insulin injections and the muscle abnormalities reported in the type 1 diabetes population. Indeed, this relationship might be considered in the context of potential cellular abnormalities in other relevant tissues, such as liver and adipose tissue.
A major clinical concern in this study is that the mitochondrial/metabolic alterations observed were in young adults with type 1 diabetes, who had self-reported moderate-to-vigorous activity levels above the American and Canadian diabetes associations’ recommendations. In addition, these changes occurred in large, proximal muscle groups without a detectable loss of capillary density. Clearly, larger cohort studies are needed to assess the impact of type 1 diabetes in those who are sedentary, as well as those who meet or exceed physical activity recommendations, to determine whether an optimal level of physical activity exists that is sufficient to restore/overcome the metabolic abnormalities that characterise the muscle of those with type 1 diabetes.
The authors are extremely grateful and thankful to all the participants for their contributions (time and tissue) to this study. The authors also thank A. N. Belcastro (School of Kinesiology and Health Science, York University, Toronto, ON, Canada) for access to his human clinical laboratory. The authors express their appreciation to J. Hanson (Executive Director; Connected in Motion, ON, Canada) for her assistance with participant recruitment, K. Mackett (McMaster University, Hamilton, ON, Canada) for her assistance in data analysis and G. R. Steinberg (Department of Medicine, McMaster University, Hamilton, ON, Canada) for generously providing some of the antibodies used in this study.
CMFM, MCH, MAT, TJH and CGRP designed the experiments. CMFM, TJH and CGRP wrote the manuscript. CMFM, MCH, SVR, NEV, CL and CM performed experiments. CMFM, MCH, SVR, NEV, CL, FAR, CM, MAT, MPK, RL, TJH and CGRP analysed and interpreted data. All authors edited the manuscript. All authors provided final approval of the version to be published. All people designated as authors qualify for authorship, and all those who qualify for authorship are listed. TJH and CGRP are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of data and the accuracy of the data analysis.
This project was funded by the Natural Sciences and Engineering Research Council of Canada (NSERC; grant 298738-2013 to TJH and grant 436138-2013 to CGRP) and the James H. Cummings Foundation (to CGRP). CMFM and MCH were recipients of NSERC graduate scholarships. CMFM was also a recipient of the DeGroote Doctoral Scholarship of Excellence. SVR was a recipient of the Ontario Graduate Scholarship.
Duality of interest
The authors declare that there is no duality of interest associated with this manuscript.
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