The effects of muscle contractions on the profile of postcontraction resting intracellular Ca2+ ([Ca2+]i) accumulation in Type 1 diabetes are unclear. We tested the hypothesis that, following repeated bouts of muscle contractions, the rise in resting [Ca2+]i evident in healthy rats would be increased in diabetic rats and that these changes would be associated with a decreased cytoplasmic Ca2+-buffering capacity. Adult male Wistar rats were divided randomly into diabetic (DIA; streptozotocin, ip) and healthy control (CONT) groups. Four weeks later, animals were anesthetized and spinotrapezius muscle contractions (10 sets of 50 contractions) were elicited by electrical stimulation (100 Hz). Ca2+ imaging was achieved using Fura-2 AM in the spinotrapezius muscle in vivo (i.e., circulation intact). The ratio (340/380 nm) was determined from fluorescence images following each set of contractions for estimation of [Ca2+]i. Also, muscle Ca2+ buffering was studied in individual myocytes microinjected with 2 mM Ca2+ solution. After muscle contractions, resting [Ca2+]i in DIA increased earlier and more rapidly than in CONT (P < 0.05 vs. precontraction). Peak [Ca2+]i in response to the Ca2+ injection was significantly higher in CONT (25.8 ± 6.0% above baseline) than DIA (10.2 ± 1.1% above baseline). Subsequently, CONT [Ca2+]i decreased rapidly (<15 s) to plateau 9–10% above baseline, whereas DIA remained elevated throughout the 60-s measurement window. No differences in SERCA1 and SERCA2 (Ca2+ uptake) protein levels were evident between CONT and DIA, whereas ryanodine receptor (Ca2+ release) protein level and mitochondrial oxidative enzyme activity (succinate dehydrogenase) were decreased in DIA (P < 0.05). In conclusion, diabetes impairs resting [Ca2+]i homeostasis following muscle contractions. Markedly different responses to Ca2+ injection in DIA vs. CONT suggest fundamentally deranged Ca2+ handling.
- calcium buffering
- Fura-2 AM
- muscle contraction
in diabetes mellitus (dia), force production in skeletal muscle is impaired, while fatigability and muscle fragility are increased (4, 5, 14, 44). Maintenance of Ca2+ homeostasis across repeated muscle contractions is requisite for optimal contractile function (9). It is generally accepted that limitations of sarcoplasmic reticulum (SR), Ca2+ handling (i.e., Ca2+ release-uptake) can represent an important component in the etiology of muscle fatigue (1, 2, 22, 50). Although those mechanisms underlying impaired SR Ca2+ regulatory functions in fatigued skeletal muscle have not been completely elucidated, there is evidence that metabolic perturbations (e.g., ADP, H+, Pi, reactive oxygen species) within the cell affect intracellular Ca2+ ([Ca2+]i) (17, 23). The observed rise in resting [Ca2+]i during muscle fatigue has been observed in isolated toad (43) and mouse single muscle fiber preparations using a low-frequency fatigue protocol (12). Our recent study has also demonstrated that the resting level of [Ca2+]i is increased after tetanic contractions in the in vivo (i.e., circulation intact) muscle preparation model (47, 48). [Ca2+]i accumulation may result from attenuation of SR Ca2+ uptake and/or leak from the SR and/or extracellular space.
The effects of continuous muscle contractions on [Ca2+]i accumulation kinetics are unclear in diabetes. Furthermore, there are no direct measurements of Ca2+ handling in diabetic in vivo models. It is well known that the Ca2+ itself causes Ca2+ release from SR (20), and we, therefore, employed direct Ca2+ injections in vivo in our model and examined the [Ca2+]i profiles in diabetic skeletal muscle. We hypothesized that, following repeated bouts of muscle contractions, the rise in [Ca2+]i evident in healthy rats would be magnified in muscles of diabetic rats and further that these changes would be associated with attenuation of the Ca2+-handling system within the in vivo environment.
Male Wistar rats (n = 43, 10 wk of age; Japan SLC, Shizuoka, Japan) were used in this study. Rats were maintained on a 12:12-h light-dark cycle and received food and water ad libitum. Rats were divided into two groups: healthy control (CONT) and diabetic (DIA) rats. Rats were anesthetized using isoflurane and given intraperitoneal injection of 45 mg/kg body wt of streptozotocin (STZ; S0130, Sigma-Aldrich St. Louis, MO) prepared fresh in saline solution. CONT animals were injected with the saline solution. Urine glucose levels of rats were measured (New Uriesu Ga, Terumo, Japan) 2 days after STZ injection with the onset of diabetes raising glucose concentrations above 500 mg/dl. These measurements of urine glucose were continued each week for 4 wk. At experiment completion, blood was collected from a tail vein puncture to confirm that the blood glucose level exceeded 300 mg/dl. All experiments were conducted under the guidelines established by the Physiological Society of Japan and were approved by University of Electro-Communications Institutional Animal Care and Use Committee. The rats were anesthetized using pentobarbital sodium (60 mg/kg ip), and supplemental doses of anesthesia were administered as needed. At the end of experimental protocols, animals were killed by pentobarbital sodium overdose.
All experimental techniques, including the spinotrapezius muscle preparation, were performed, as described previously (47). Briefly, the right spinotrapezius muscle was gently exteriorized with minimal blood loss and tissue/microcirculatory damage and attached to a wire horseshoe around the caudal periphery by six equidistant sutures placed around the muscle perimeter. For the contraction protocols, electrodes were placed on the dorsal spinotrapezius surface along the caudal periphery, facilitating whole muscle contractions. The exposed muscle tissue was kept moist by superfusing with warmed Krebs-Henseleit buffer (KHB; 132 mM NaCl, 4.7 mM KCl, 21.8 mM NaHCO3, 2 mM MgSO4, and 2 mM CaCl2) equilibrated with 95% N2-5% CO2 and adjusted to pH 7.4, at 37°C. The fluorescent Ca2+ indicator Fura-2 AM (5 mM; Dojindo Laboratories, Kumamoto, Japan) was dissolved in DMSO and Pluronic F-127 and dispersed into KHB solution at a final concentration of 20 μM. The muscles were incubated in Fura-2 AM/KHB solution for 30–60 min on a 37°C hotplate. After incubation, muscles were rinsed with dye-free KHB solution to remove nonloaded Fura-2.
The spinotrapezius muscles loaded with Fura-2 AM were mounted on the 37°C glass hotplate (Kitazato Supply, Shizuoka, Japan) and observed by fluorescence microscopy using a 10× objective lens (0.30 numerical aperture; Nikon, Tokyo, Japan). After ensuring that the spinotrapezius muscle was not grossly damaged and supported robust capillary blood flow, a sampling area (∼880 × 663 μm) was selected using branching vessels as landmarks, and bright-field images were captured. Thereafter, 340-nm and 380-nm wavelength excitation light was delivered using a Xenon lamp equipped with appropriate fluorescent filters, and pairs of fluorescence images were captured through the 510-nm emission wavelength filter for ratiometry.
Repeated bouts of muscle contractions: experiment 1.
Spinotrapezius muscles from CONT and DIA rats were stimulated to contract, as described in detail previously (47). The time course of [Ca2+]i change was observed after each of 10 discrete sets of isometric tetanic muscle contractions. Each set consisted of the muscle being stimulated tetanically at resting spinotrapezius sarcomere length (100 Hz, 5–8 V, stimulus duration 700 ms, 2.6- to 2.8-μm sarcomere length) every 3 s for 2.5 min (i.e., 50 contractions). Pairs of fluorescence images were captured precontraction, and after each set of contractions, as well as at the end of the 5-min between-set recovery (immediately before initiation of the subsequent set of contractions). Images were captured by a charge-coupled device digital camera (DP70; Olympus, Tokyo, Japan) using image-capture software (DP Control; Olympus, Japan). After selecting an appropriate region of interest, which included multiple muscle fibers, the spinotrapezius fluorescence was observed. Images were converted to 340/380 ratio (F340/F380: R) image by ImageJ software [National Institutes of Health (NIH), Bethesda, MD], and the ratio image data, indicating Ca2+ levels, were averaged over the whole area sampled. The fluorescence intensity of serial ratio images was normalized to the starting point (i.e., precondition, R0) of each experiment (R/R0).
Microinjection with high Ca2+ solution: experiment 2.
Capillary micropipettes were generated with a tip of 10 μm in diameter, which was achieved by custom grinding and inserted into the selected single muscle fiber using a micromanipulator precision-controlled advancer (MMO-220A; Narishige, Tokyo, Japan). Subsequently, using a microinjector (IM300; Narishige, Japan), a single muscle fiber was microinjected with 2 mM Ca2+ solution by microinjection at 35 psi (24,000 Pa) for 1 s. Criteria for successful injection included an unchanged [Ca2+]i in adjacent muscle fibers. Fluorescence images were captured by a high-sensitivity charge-coupled device digital camera (ORCA-Flash2.8; Hamamatsu Photonics, Hamamatsu, Japan) using image-capture software (NIS-Elements Advanced Research; Nikon, Tokyo, Japan) at a 1-s exposure for 60 s after injection. Pilot studies confirmed that insertion of the capillary micropipette itself did not induce any [Ca2+]i change.
At the end of the experiments, the spinotrapezius muscle was resected under anesthesia. The tissue blocks were frozen rapidly in isopentane cooled in liquid nitrogen. Serial sections of 10 μm were made with a cryostat (CM1510; Leica, Wetzlar, Germany) at −20°C and mounted on polylysine-coated slides. Whole sections were stained for hematoxylin-and-eosin, succinate dehydrogenase (SDH), and slow-type myosin heavy chain (MHC). The SDH activities in individual muscle fibers in histological sections were examined as an index of mitochondrial volume density. Sections were incubated at 37°C for 45 min in a medium consisting of 0.2 M sodium phosphate buffer pH 7.5, containing 0.2 M sodium succinate and 1.2 mM nitroblue tetrazolium. Mouse-monoclonal slow MHC antibody (diluted 1:40 in PBS) supplied by Novocastrate Laboratories (Leica Biosystems) was used to identify slow-twitch fibers. The sections were allowed to warm to room temperature and incubated in PBS (pH 7.5) at 25°C before further incubation with the primary antibody in a humidified box overnight at 4°C. Vectastain ABC kit (PK-6102; Vector Laboratories, Burlingame, CA) was used to reveal the immunohistochemical reaction, according to the manufacturer's instructions. Then, DAB peroxidase substrate kit (SK-4100; Vector Laboratories) was used as a chromogenic reaction.
In photographs of serial sections, slow MHC-stained muscle fibers were identified, and nonstained fibers were considered to be fast twitch. SDH activity was recorded using a camera (E800, 0.30 numerical aperture; Nikon) at 10× magnification and analyzed subsequently in ImageJ (NIH, Bethesda, MD). The cross-sectional areas and SDH activities were measured by tracing fiber outlines of ∼220 fibers from the muscle sections. The images were digitized as gray-level pictures. Each pixel was quantified as one of 256 gray levels and then automatically converted to optical density using ImageJ software.
Western Blot Analysis
Western blotting was performed to determine the protein expression levels of SR Ca2+-ATPase 1 (SERCA1), SR Ca2+-ATPase 2 (SERCA2), and ryanodine receptor (RyR) in spinotrapezius muscle from CONT and DIA rats. The spinotrapezius muscles were removed and homogenized in ice-cold lysis buffer (50 mM Tris, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 200 mM NaF, 20 mM sodium pyrophosphate, 1 mM NaVO4 mM, 1% Nonidet P-40, 10% glycerol) with Protease inhibitor cocktail (Nacalai Tesque, Kyoto, Japan) and PhosSTOP phosphatase inhibitor cocktail (Roche Applied Science, Indianapolis, IN). Homogenates were centrifuged at 14,000 rpm for 30 min at 4°C. Supernatant proteins were then quantified using BCA protein assay kit (Thermo Scientific, West Palm Beach, FL). The sample (10 μg total protein per lane) were separated on 7.5% (for SERCA1 and SERCA2) or 4% (for RyR) polyacrylamide gels for 50 min at 150 V and then transferred to Amersham Hybond-P membranes (GE Healthcare, Buckinghamshire, UK) for 60 or 100 min at 72 mA using the semidry method. After the transfer, the membranes were blocked with Blocking One or 1% skim milk (Nacalai Tesque) at room temperature for 1 h. After blocking, the membranes were then incubated with primary antibodies (anti-SERCA1 IIH11 antibody, 1:2,500; Thermo Scientific, MA3-911; anti-SERCA2 2A7-A1 antibody, 1:2,000; Thermo Scientific, MA3-919; and anti-RyR, 34C antibody, 1:5,000, Thermo Scientific, MA3-925) at 4°C for over-night. Then, the membranes were added to goat anti-mouse IgG linked to peroxidase (SC-2055; Santa Cruz Biotechnology, Santa Cruz, CA) for 1 h at room temperature. The bound antibodies were detected by Chemi-Lumi One Super Kit (Nacalai Tesque) and analyzed with ImageQuant LAS-4000 (GE Healthcare Life Sciences, Tokyo, Japan). SERCA1, SERCA2, and RyR protein levels were expressed relative to α-tubulin antibody (1:1,000, GT114; GeneTex, Irvine, CA) and normalized to the CONT samples.
All experimental data are expressed as means ± SE. All statistical analyses were performed in Prism version 4.0 (GraphPad Software, San Diego, CA). A two-way repeated-measures ANOVA and Bonferroni post hoc test were used for group differences in [Ca2+]i levels and SDH activity. A one-way repeated-measures ANOVA and Bonferroni post hoc test were used for relative force comparison in repeated bouts of muscle contractions. t-tests were used for blood glucose level, histological data, and relative protein levels. The level of significance was set at P < 0.05.
Blood glucose concentration was 98.6 ± 6.1 (range of 89 to 128) and 393.6 ± 35.3 (range of 317 to 600) mg/dl in CONT and DIA, respectively (P < 0.01). DIA rats evidenced a significant decrease in body weight at 4 wk post-STZ injection (Table 1), and these animals demonstrated a significant reduction in spinotrapezius fast-twitch fiber cross-sectional area (Fig. 1, Table 1). Also, muscle fiber type shifts from fast to slow were found in DIA (32.5 ± 2.0% to 45.1 ± 5.9%, P < 0.01). Figure 2 shows histograms of SDH activity in each muscle fiber cross-sectional area distribution. DIA decreased SDH activity significantly below that found in CONT in all fiber cross-sectional area categories, except the largest (i.e., >2,500 μm2).
After repeated bouts of muscle contractions (experiment 1), Fura-2 ratios were increased in both CONT and DIA. [Ca2+]i became significantly elevated above baseline only after six sets of contractions in CONT (Fig. 3). In marked contrast, significant [Ca2+]i accumulation in DIA occurred following the very first set of contractions. After the final contraction set (10 sets), [Ca2+]i in DIA was elevated 32.0 ± 8.7% (P < 0.01) above precontraction baseline vs. ∼10% for CONT.
As shown in Fig. 4, isometrically evoked active force in CONT decreased ∼30% within a given set of contractions, which was less than the ∼40% seen for DIA (P < 0.01). At the start of set 5, relative force in CONT was ∼90% of initial baseline compared with ∼60% for DIA, indicating a greater compromise of tension recovery after contractions in DIA. Moreover, for the final (10th) set, the muscle force of the first and last 5 contractions in DIA was significantly lower compared with CONT.
Elevations of [Ca2+]i following Ca2+ microinjections (experiment 2) were significantly higher in CONT compared with DIA (Figs. 5 and 6). The high [Ca2+]i region after the injection was larger and more pronounced in CONT than DIA (Fig. 5, B vs. F). The [Ca2+]i peak value of CONT occurred 6–8 s after the injection. In contrast, the [Ca2+]I peak value of DIA was far lower and was evidenced earlier than CONT (i.e., 4–6 s after the injection, Fig. 6). In CONT muscles following the [Ca2+]i peak, [Ca2+]i fell rapidly for 10–15 s to stabilize at 8–10% above the initial baseline. On the other hand, [Ca2+]i in DIA muscle fibers rose ∼8–10% above baseline and did not decrease subsequently (i.e., to the 60-s observation mark, Fig. 6).
Figure 7 shows the sarcoplasmic reticulum-related proteins (SERCA1, SERCA2, and RyR). No differences (P > 0.05) in SERCA1 and SERCA2 expression levels were noted between CONT and DIA (Fig. 7, A and B), whereas RyR content was decreased ∼50% (P < 0.05) in spinotrapezius muscles from DIA compared with CONT (Fig. 7C).
This investigation represents the first in vivo imaging of [Ca2+]i dynamics associated with (i.e., following) repeated tetanic contractions and direct single-fiber Ca2+ loading in skeletal muscles of diabetic rats. The principal original findings were that [Ca2+]i failed to return to baseline levels earlier (i.e., first set) and increased more than twice as rapidly in DIA than in CONT muscles following contractions. Moreover, a pronounced dysfunction of Ca2+ release-reuptake dynamics was identified in DIA myocytes. It is likely that these effects are linked mechanistically to the reduced muscle contractility and impaired exercise tolerance from which diabetic patients suffer.
Effects of Diabetes on Morphological Properties of Spinotrapezius Muscle
The rat spinotrapezius muscle possesses the following important characteristics that make it an excellent candidate for intravital microscopy studies: 1) it can be exteriorized and transilluminated without disruption of the nervous or primary vascular supplies (6, 34, 35, 40); 2) it comprises both principal fiber types (41% Type I; and 59% Type II); and 3) its oxidative capacity approximates that found in the untrained human quadriceps (16, 38). Until the present investigation, the impact of the DIA condition on the muscle fiber composition and oxidative enzyme activity was unknown. We found that, in addition to DIA causing a pronounced atrophy in the fast-twitch fibers, there was a shift toward a greater slow-twitch fiber composition (Fig. 1, Table 1). This result was consonant with previous studies showing that the STZ-induced diabetic rat endures a fast-twitch, fiber-specific atrophy of hindlimb muscles (5, 14, 39). Also, DIA induced a profound decrease of SDH activity compared with CONT (Fig. 2), as predicted from hindlimb locomotory muscle studies (30, 36).
Effects of Muscle Contractions on [Ca2+]i Accumulation
Muscle contraction is evoked by the transient elevation of [Ca2+]i, which under normal conditions, returns almost immediately to basal resting levels. In fact, previous studies indicate that [Ca2+]i accumulation may not occur following shorter bouts of isometric tetanic contractions (7). On the other hand, longer-lasting bouts of fatiguing muscle contractions do elicit muscle [Ca2+]i accumulation (11, 27, 28, 47). In the present investigation, elevation of [Ca2+]i postcontractions (i.e., at rest) in DIA was greater and occurred after fewer bouts of contractions compared with that in CONT. However, it is unlikely that these responses are directly responsible for the DIA tension deficits. Specifically, Allen et al. (2) showed the beneficial effects of caffeine, which enhanced/restored force development in repetitive tetanic stimulation-fatigued isolated single muscle fibers via increased SR Ca2+ release. They proposed that it was the impairment of Ca2+ release from the SR, which constituted a key factor in the fatigue development (3).
Although the protein content per se did not change in DIA, we speculate that the most likely source of the elevated [Ca2+]i was SR-released Ca2+, and its accumulation reflects a progressive inability for the SR to recover during each resting period. Because Ca2+ uptake by the SR is actively transported, it is associated with an ATP supply level. Kindig et al. (35) found that peripheral circulatory function of STZ-diabetic rats is decreased. The reduction in metabolic potential induced by this ischemia and the associated hypoxic/anoxic environment may be responsible, in part, for the observed accumulation of [Ca2+]i in DIA. Previous studies demonstrated that ischemia-induced stress increased [Ca2+]i in skeletal muscle (32).
The importance of prolonged elevation of resting [Ca2+]i after contractions likely relates to the muscle weakness and/or activation of proteolytic pathways (12, 55, 56). One plausible mechanistic link between the proteolysis of cytoskeletal proteins and elevated [Ca2+]i is activation of Ca2+-activated neutral proteases (calpains) (29, 55). The repeated eccentric-contraction protocol induces muscle damage causing calpain-3 autolysis and elevated [Ca2+]i immediately after contractions (48). Therefore, the lack of [Ca2+]i homeostasis in diabetes may be associated with the diabetogenic fragility, such as muscle atrophy or damage.
Effects of Ca2+ Injection on [Ca2+]i Kinetics
As described above, the failure of diabetes-related Ca2+ homeostasis may be associated with the compromised SR function. In previous studies using cardiac muscle, decreased SR Ca2+ transport and SR Ca2+ content have been detected in animal diabetic models (13, 19). However, there are conflicting reports on the capability of the SR to function in diabetic skeletal muscle. Dhalla and colleagues have reported that SR Ca2+ transport activity actually increases in skeletal muscle during the development of diabetes (25, 53). On the other hand, and as noted above, Racz et al. (41) recently reported that level of SR Ca2+-ATPase protein declined in STZ diabetic rats. With respect to an in vivo experimental model (circulation intact), we believe that the present investigation is the first to use Ca2+ solution microinjection directly into the cytoplasm to determine the impact of DIA on the SR system. Figures 5 and 6 provide strong evidence that diabetes impairs Ca2+ handling in skeletal muscle.
In cardiac muscle, the Ca2+-induced Ca2+ release (CICR) is considered to be the physiological mechanism responsible for cardiac muscle contraction (21, 46, 51). In skeletal muscle, however, the primary mechanism of physiological Ca2+ release is not CICR, but rather interaction between the voltage sensor of the t-tubule membrane, the dihydropyridine receptor and the RyR (20, 45). RyR1 is the primary isoform in mammalian skeletal muscle (49). As all types of RyR (i.e., RyR1, RyR2, and RyR3) show CICR activity (20), the present investigation can quantify dynamically the Ca2+ release by the SR system. Our data reveal that diabetic rats significantly decreased peak [Ca2+]i following Ca2+ injection. Correspondingly, attenuation of the Ca2+ release response in DIA was accompanied by reduction of RyR protein level in the spinotrapezius muscle (Fig. 7). In cardiomyocytes, previous studies indicated that there was a lower density of RyR and a quantitative relationship between the altered contraction-relaxation cycle and reduced Ca2+ release (31, 42, 52). In fact, Venetucci et al. (54) reported a lower transient amplitude of [Ca2+]i consistent with a decreased SR calcium content.
An important role of the SR system is Ca2+ reuptake to maintain [Ca2+]i at extremely low levels. In the present investigation, the DIA myocytes had almost no discernible [Ca2+]i recovery after Ca2+ injection. On the other hand, no significant difference was found in the protein expression level of the SR Ca2+ pump (SERCA1 and SERCA2) between CONT and DIA. Taira et al. (53) reported that Ca2+ transport activities are enhanced in the skeletal muscle of STZ rats using in vitro biochemical assay. They suggested that enhanced Ca2+ transport activities may be related to the many metabolic changes arising from the insulin deficiency in the STZ model. For example, the high level of circulating catecholamines induced by diabetes may be associated with an increased Ca2+ pump function (26). Recently, Funai et al. (24) showed impairment in SR Ca2+ transport activity without changes in SERCA1 expression in fatty acid synthase knockout mice. Thus, in the present investigation, altered fatty acid synthetase in DIA may have altered SERCA1 activity independent of protein levels and, as a result, impacted Ca2+ dynamics in DIA following the Ca2+ injection.
Another putative source of Ca2+ uptake is the mitochondria, and it is known that Ca2+ release from the SR through RyR also promotes Ca2+ uptake into neighboring mitochondria (15, 57). In skeletal muscle fibers, the mitochondrial reticulum is located adjacent to the SR and close to Ca2+ release units (8), indicating that mitochondria and SR function in a complementary fashion to regulate [Ca2+]i dynamics. This notion is supported by the observation that mitochondrial poisoning by carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone decreases the amplitude of the Ca2+ transient associated with excitation-contraction coupling in isolated skeletal muscle fibers (10). Moreover, Eisner et al. (18) found that temporal loss of connectivity of the mitochondria in skeletal myotubes compromised mitochondrial Ca2+ buffer capacity. The lowered SDH activity in DIA indicates a reduced quantity and/or quality of mitochondria that may, consequently, contribute to the impaired [Ca2+]i regulation demonstrated herein (Figs. 3, 5, 6).
Perspectives and Significance
The present investigation demonstrates profound impairments in [Ca2+]i homeostasis following contractions in diabetic skeletal muscle. Although the SERCA protein content was unchanged in DIA, our in vivo data indicate that diabetes-incurred dysfunction of both Ca2+ release and reuptake mechanisms [involving RyRs and mitochondrial (dys)function] are likely to be responsible for this behavior and may account, in large part, for the decline of the muscle contractile function by diabetes. It is also known that elevated [Ca2+]i activates calpains and may, therefore, be involved in the chronic morphological changes (myocyte atrophy, loss of mitochondrial capacity) characteristic of this condition.
The present investigation was performed using adult STZ-induced diabetic animals. Hawke and colleagues (33) demonstrated that direct action of STZ (i.e., independent of hyperglycemia) evokes changes in muscle fiber growth (33) and muscle twitch characteristics (37). In particular, they suggested the possibility that STZ directly influences Ca2+ handling, such that it will be important to study alternative models of Type 1 diabetes mellitus to clarify the relationship between the hyperglycemic state and Ca2+ handling.
This study was supported in part by Grant-in-Aid for Scientific Research from Japan Society for the Promotion of Science (no. 22300221 and 2365043) and Yamaha Motor Foundation for Sports.
No conflicts of interest, financial or otherwise, are declared by the authors.
Author contributions: H.E. and Y.K. conception and design of research; H.E., Y.T., T.S., T.I., and Y.K. performed experiments; H.E., Y.T., T.S., T.I., T.N., and Y.K. analyzed data; H.E., Y.T., T.S., T.I., D.C.P., and Y.K. interpreted results of experiments; H.E., D.C.P., and Y.K. prepared figures; H.E., D.C.P., and Y.K. drafted manuscript; H.E., T.N., D.C.P., and Y.K. approved final version of manuscript; D.C.P. and Y.K. edited and revised manuscript.
We gratefully acknowledge Dr. Mizuki Sudo (Fukuoka Univ.) for technical assistance on microscopy and fluorescence analysis.
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