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Am J Physiol Regul Integr Comp Physiol. Mar 15, 2012; 302(6): R712–R719.
Published online Dec 7, 2011. doi:  10.1152/ajpregu.00229.2011
PMCID: PMC3774485

Measuring mitochondrial respiration in intact single muscle fibers

Abstract

Measurement of mitochondrial function in skeletal muscle is a vital tool for understanding regulation of cellular bioenergetics. Currently, a number of different experimental approaches are employed to quantify mitochondrial function, with each involving either mechanically or chemically induced disruption of cellular membranes. Here, we describe a novel approach that allows for the quantification of substrate-induced mitochondria-driven oxygen consumption in intact single skeletal muscle fibers isolated from adult mice. Specifically, we isolated intact muscle fibers from the flexor digitorum brevis muscle and placed the fibers in culture conditions overnight. We then quantified oxygen consumption rates using a highly sensitive microplate format. Peak oxygen consumption rates were significantly increased by 3.4-fold and 2.9-fold by simultaneous stimulation with the uncoupling agent, carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP), and/or pyruvate or palmitate exposure, respectively. However, when calculating the total oxygen consumed over the entire treatment, palmitate exposure resulted in significantly more oxygen consumption compared with pyruvate. Further, as proof of principle for the procedure, we isolated fibers from the mdx mouse model, which has known mitochondrial deficits. We found significant reductions in initial and peak oxygen consumption of 51% and 61% compared with fibers isolated from the wild-type (WT) animals, respectively. In addition, we determined that fibers isolated from mdx mice exhibited less total oxygen consumption in response to the FCCP + pyruvate stimulation compared with the WT mice. This novel approach allows the user to make mitochondria-specific measures in a nondisrupted muscle fiber that has been isolated from a whole muscle.

Keywords: oxygen, metabolism

the understanding of mitochondrial function in skeletal muscle is a critical component for delineating mechanisms behind muscular adaptation and their contribution to various chronic health conditions. It is now well established that mitochondrial function contains some plasticity, and deficiencies in function are critical to the pathogenic progression of numerous muscle diseases.

Several approaches have been used to assay mitochondrial function in muscle, and each has limitations and benefits. For example, it has become common to use measurements of single mitochondrial enzymes as a surrogate to measuring mitochondrial function. Unfortunately, because of the complexity of mitochondrial activity, these measures do not provide a complete physiological readout of mitochondrial function but instead only provide information about the activity of that specific enzyme. It is assumed that the changes in single enzyme activity frequently correlate with changes in overall mitochondrial function. However, there are often circumstances when not all mitochondrial enzyme activity is altered to the same magnitude under a chosen condition, making interpretation of the data difficult.

Measurements of mitochondrial respiration provide a more appropriate indication of the function of mitochondria. Historically, these measures have been made using amperometric oxygen electrodes (i.e., the Clark electrode) to determine changes in oxygen tension within a buffer. With this approach, mitochondrial respiration studies in skeletal muscle have been typically conducted in isolated mitochondria or in permeabilized muscle fiber bundle preparations. The isolated mitochondrial approach requires fresh muscle samples that are mechanically homogenized and subsequently exposed to a differential centrifugation approach, resulting in a reasonably pure mitochondrial fraction (12). The isolation yields a high mitochondrial purity, but evidence suggests that the isolation procedure can result in the loss of specific complexes within the electron transport chain (27). Furthermore, recent evidence has documented that this approach can disrupt or even overestimate mitochondrial function (26).

Because of limitations with isolated mitochondria preparations, permeabilized fiber bundles are often used to measure oxygen consumption of mitochondria. Fiber bundles are mechanically dissected from the whole muscle and chemically treated to permeabilize the sarcolemma without damaging other organelle membranes (20, 30). This approach allows investigators to measure mitochondrial function in the context of the whole muscle fiber with minimal mechanical disruption, while maintaining control over the intercellular milieu. A disadvantage is that disruption of the muscle fiber membrane necessitates measures be made immediately upon tissue removal. In addition, in this technique, because the cytoplasm and external media become equilibrated, any substances within the equilibration media could affect mitochondrial function. Thus, great care must be taken to ensure proper media formulation. Both of these considerations place time and experimental constraints on the experiment, which affects the throughput of this approach. However, some groups have shown that prior to permeabilzation, it is possible to cyropreserve isolated fiber bundles for subsequent measures with minimal effect on mitochondrial function (19, 34). This approach would allow for increased sample collection without the need for immediate processing, but it is unclear why this procedure has not resulted in more utilization.

Mitochondrial function has also been assessed in intact nondisrupted tissue/cellular preparations, and important information has been gained. The main benefit of this approach is that it allows mitochondrial function to be assessed in its most native state. The main drawback of whole muscle experiments with classic amperometric oxygen probes is the diffusion limitation of oxygen in the tissue and bath. More recently, miniaturized self-referencing or amperometric electrodes have proved particularly useful in overcoming diffusion limitations by allowing the assay of oxygen metabolism in single cultured cell preparations (23); however, the techniques are highly demanding and thus result in lower throughput. For example, Elzinga and van der Laarse (8) eloquently demonstrated methodology using oxygen electrodes and isolated single fibers from frog muscle, where they demonstrated increasing oxygen consumption as a function of contraction frequency (8). It was determined that the maximum rate of oxygen consumption by the fibers was related to activity levels of the mitochondrial protein succinate dehydrogenase (35).

Recently, systems have been developed to make measurements of mitochondrial respiration in the same small volumes within a multiwell microplate format coupled with substantially enhanced sensitivity compared with standard amperometric oxygen probes (25). With this approach, measurement sensitivity in both isolated mitochondria and primary cell cultures has been greatly enhanced, allowing for increased throughput and enabling treatments and replicates to be conducted rapidly. Here, we describe the use of this approach to assay single intact skeletal muscle cells.

We have adapted the use of intact individual muscle fibers isolated from a whole muscle to measure mitochondrial respiration. This approach allows for similar measures to be made, as previously documented; however, because the fiber membranes are not disrupted, it allows the user the ability to place the fibers in culture, removing the need to make immediate measures after the isolation procedure. In addition, this approach assesses the ability of the mitochondria within the fibers to consume oxygen at a physiological temperature. Finally, after the respiration experiments, the user can use the cells for immunohistochemical techniques or isolate the cells for procedures such as Western blot analyses, quantitative PCR, or activity assays.

MATERIALS AND METHODS

Animals.

For the first series of studies, 10- to 16-wk-old male C57/BL6 mice were used. In our second studies, the murine model of Duchenne's muscular dystrophy (C57BL/10 ScSn-mdx) (mdx) and their control counterparts (C57BL/10 ScSn), hereafter referred to as wild-type (WT) mice, were used at 5 mo of age. All experiments were approved by the University of Maryland Animal Care and Use Committee.

Chemicals.

All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless otherwise indicated.

Single fiber isolation.

After euthanasia (CO2 inhalation), flexor digitorum brevis (FDB) muscles were harvested bilaterally from C57/BL6, mdx, or WT mice. Single skeletal muscle fibers were enzymatically isolated from the FDB muscle. An example microscope bright-field image of the isolated intact single fibers and a high-magnification image of an individual fiber stained with MitoTracker Green/DAPI (blue) are shown in Fig. 1. The isolation procedure was modified from work previously published with this method in muscle from rats and mice (25, 22, 28). In brief, surgically excised FDB muscles were incubated in dissociation media (DM) containing DMEM (Invitrogen, Carlsbad, CA), gentamycin ( 50 μg/ml), FBS (2%, no. 30–2020; American Type Culture Collection, Rockville, MD), and collagenase A (4 mg/ml; Roche, Mannheim, Germany). Two FDB muscles were placed in a 35-mm disposable culture dish with 4 ml of DM and then in an incubator (37°C, 5% CO2) for 1.5–2 h. Following the dissociation, muscles were placed in a new 35-mm plate with warmed media containing gentamycin and FBS but without collagenase. FDB muscles were triturated with a small bore (~1 mm) fire-polished glass transfer pipette to yield single FDB myofibers. If trituration did not yield a significant number of disassociated single fibers after 5–10 passes, the muscles were returned to DM for 15–30 min, and the process was repeated. Following trituration, large debris (nerve, undigested FDB muscle) was removed with forceps. Seahorse XF24 cell culture V7 microplates (Seahorse Bioscience, Billerica, MA) were coated with 5 μl of extracellular matrix (ECM; Sigma-Aldrich, E1270, St. Louis, MO) previously diluted 1:1 in DMEM. Following application of ECM in the wells, the lid was placed on the V7 microplate, and the plate was vigorously tapped horizontally against an open hand to spread the ECM throughout the wells. The plate was allowed to briefly air dry for 10–30 min. Following a thorough dispersion of single fibers in the 35-mm dish, 75-μl aliquots of the media were taken randomly and deposited into each well with the goal being to allow the fibers to cover ~60% confluency of the well bottom. The confluency was determined through light microscope visualization. Single myofibers were verified as being fully adherent within 5 min. The cells were placed in a 95% air-5% CO2 humidified incubator at 37°C overnight in DM media without the collagenase.

Fig. 1.
A and B: visual representation of intact single muscle fibers isolated from the flexor digitorum brevis (FDB) of adult male mice. A: bright-field image of cultured single fibers after dissociation (×10 magnification). Scale bar = 500 μm. ...

XF24 microplate-based respirometry.

Bioenergetic analyses of intact single cultured muscle fibers were performed using an XF24-3 Extracellular Flux Analyzer (Seahorse Bioscience). As previously described, the Seahorse Bioscience technology provides an innovative and more sensitive way to assess mitochondrial function compared with traditional amperometric oxygen probes, which results in real-time sensitive measures with high throughput due to the microplate format (13, 25).

Intact fibers.

The assay measurement buffer (MB) at ~37°C contained 120 mM NaCl, 3.5 mM KCl, 1.3 mM CaCl2, 0.4 mM KH2PO4, 1 mM MgCl2, 5 mM HEPES (pH 7.4) supplemented with 2.5 mM d-glucose (Sigma G7528) and 0.5 mM l-carnitine (Sigma CO158). Prior to measurements, prewarmed MB was gently added to the fibers, and the fibers were placed in an unbuffered, humidified incubator at 37°C for 2 h to allow temperature and pH equilibration. Because the Flux Analyzer performs direct injections of substrates to the media in the wells, it is absolutely critical that the investigator know the initial volume of media in each well prior to the addition of the MB. Thus, 1,000 μl of MB was removed from each well, and 600 μl of MB was added per well prior to placing the plate in the analyzer, resulting in a final volume of 675 μl. Each plate contains specific temperature control wells, which have no fibers in the wells. These wells had 675 μl of MB added to each well. Following this incubation period, the calibration plate was loaded onto the instrument, and calibration was initiated. Following calibration, the plate containing the fibers replaced the calibration plate in the instrument to begin the experimental run. After an equilibration step, basal oxygen consumption rates (OCR, pmol/min) and extracellular acidification rates (ECAR, milli-pH/min) were recorded using 3-min mix, 2-min wait, and 3-min measure (looped 3 times) cycles prior to injection of substrate plus carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP; Sigma C2920) to induce maximal oxygen consumption. Three more measurement loops were recorded following FCCP injection and prior to injection of antimycin A (inhibitor of mitochondrial respiration, Sigma A8674). Two measurement loops were recorded after the injection of antimycin A. The drugs prepared in MB (75 μl volumes) were preloaded and then sequentially injected as indicated through ports in the XF24 calibration cartridge to final concentrations of 400 nM FCCP, substrates supplied were either sodium pyruvate (10 mM; Sigma P8574) or albumin (Roche, Indianapolis, IN, 03117405001) conjugated to sodium palmitate (Sigma P9767; 200 μM), and antimycin A (1 μM). Upon completion of the runs, all cells were scraped into Mueller buffer [50 mM HEPES (pH 7.4), 0.1% Triton X-100, 4 mM EGTA, 10 mM EDTA, 15 mM Na4P2O7·H2O, 100 mM β-glycerophosphate, 25 mM NaF, 50 μg/ml leupeptin, 50 μg/ml pepstatin, 40 μg/ml aprotinin, 5 mM Na3VO4, and 1 mM PMSF] to lyse the cells to normalize the data to total protein (32). Unfortunately, the amount of total protein detected in each well was below the minimal detection limits of the assay (BCA assay; Pierce, Rockford, IL), and thus, we were unable to use this as a normalization approach.

Permeabilized fibers.

For these experiments, we followed the previously developed protocol with a few minor changes (20). We isolated the fibers as previously described above, and the fibers were allowed to adhere overnight to ECM-coated plates. The following morning, the fibers were placed in buffer A (10 mM Ca-EGTA buffer, 2.77 mM of CaK2EGTA + 7.23 mM of K2EGTA, free concentration of calcium, 0.1 μM; imidazole, 20 mM; taurine, 20 mM; K-MES, 49 mM; K2HPO4, 3 mM; MgCl2, 9.5 mM; ATP, 5.7 mM; phosphocreatine, 15 mM; leupeptin, 1 μM; at pH 7.1, adjusted with 5 N KOH) at room temperature for 5 min. Buffer A was replaced with buffer A containing saponin (50 μg/ml) and allowed to incubate for another 30 min. The buffer was then changed to buffer B [EGTA, 0.5 mM (0.19 g/l); MgCl2·6H2O 3 mM (0.61 g/l); taurine, 20 mM (2.502 g/l); KH2PO4, 10 mM (1.361 g/l); HEPES, 20 mM (4.77 g/l); BSA, 1 g/l; potassium-lactobionate, 60 mM (120 ml of 0.5 M K-lactobionate stock solution/l); mannitol, 110 mM (20.04 g/l); dithiothreitol, 0.3 mM (0.046 g/l); at pH 7.1, adjusted with 5 N KOH] and placed in the Extracellular Flux Analyzer for specific measures.

Statistics.

All experiments were conducted three to five times with at least five replicates per experiment. All data are expressed as means ± SE. Statistical significance was determined using a Student's t-test for comparing FCCP-stimulated OCR rates compared with basal OCR rates, area under the curve (AUC) OCR measures, and for comparison across time points of OCR between the WT and mdx fibers. A two-way ANOVA was utilized for spare respiratory capacity measures between the WT and mdx fibers (see Fig. 3D). With the ANOVA, when an interaction was found, the test was followed by a Holm-Sidak post hoc test. A P value of <0.05 was considered significant.

Fig. 3.
A–D: basal and stimulated OCR was significantly lower in the fibers from mdx mice compared with the wild-type (WT) mice. A: Initial and FCCP + Pyr-stimulated peak OCR was significantly lower in the mdx fibers (gray circles) compared with the WT ...

RESULTS

Pyruvate- and palmitate-induced respiration.

In the first set of experiments, we isolated intact single fibers from 10- to 16-wk-old male C57/BL6 mice. After the fibers were dissociated, we plated the fibers on ECM-coated XF24 V7 microplates overnight. The next day, the intact single fibers were removed from the incubator, the media were removed and replaced with MB containing low glucose concentrations. The fibers were returned to a 37°C non-CO2-buffered incubator for 2 h. The fibers were then placed in the XF24-3 analyzer for analysis. In these experiments, we sought to determine whether respiration could be stimulated in intact muscle fibers using commonly used substrates, sodium pyruvate, and albumin-conjugated palmitate. Because both substrates are transported across the sarcolemma, we supplied the substrates in excess simultaneously with the mitochondrial uncoupler FCCP, to the MB, as indicated in Fig. 2. Under basal conditions, the energetic demand of the fibers is low; thus, to enhance substrate utilization by the mitochondria, we provided the cells with FCCP. We found that both substrates resulted in significant increases (P < 0.001) of ~3-fold in oxygen consumption by the single fibers (Fig. 2A). To confirm that the increase in oxygen consumption was specific to the mitochondria, we next added antimycin A, an inhibitor of complex III of the electron transport chain. As can be seen at the point indicated in Fig. 2A, respiration was returned to baseline after antimycin treatment, confirming that the increase in oxygen consumption, mediated by the substrate exposure, was due to increased oxygen consumption by the mitochondria. We found no difference in the ECAR when the intact single fibers were treated with either pyruvate or palmitate (Fig. 2B). ECAR is an indirect measure of glycolytic activation, measured as milli-pH/min, with the assumption that in the cultured media any increase in acidification of the media driven by accretion of protons is derived by lactate (25). Thus, any increases in ECAR indicate an increased acidification of the medium potentially due to enhanced lactate production. There was no significant difference in the peak oxygen consumption induced by either substrate; however, when considering the total amount of oxygen consumed as calculated by the area under the curve (AUC, Fig. 2C), more total oxygen was consumed when the fibers were treated with palmitate.

Fig. 2.
AC: basal and stimulated oxygen consumption rates (OCR) of cultured adult intact single muscle fibers. A: initial OCR was measured in the single fibers, followed by peak OCR, which was induced (first arrow) by direct exposure to FCCP (400 nM) ...

To determine whether the isolated single fibers could be used in the extracellular flux analyzer in a similar approach, as previously described for the permeabilized fiber bundles, we conducted experiments in which we treated the isolated fibers with saponin to permeabilize the sarcolemma. Saponin is a cholesterol-specific detergent that in muscle cells will selectively permeabilize the sarcolemma without damaging the mitochondria (20, 30). Unfortunately, we found the permeabilized single-fiber approach is not compatible with the Extracellular Flux Analyzer due to a temperature sensitivity issue. Specifically, unlike our results in the intact fibers, we were unable to reliably measure substrate or FCCP-driven increases in OCR, when the fibers were permeabilized (data not shown). When we placed the permeabilized fibers in the Extracellular Flux Analyzer and exposed the fibers to 10 mM glutamate and 5 mM malate, we found no changes in OCR even if we injected a saturating dose of 2 mM ADP, which should induce a maximal State 3 respiration (20). We also were unable to induce an FCCP response in these permeabilized fibers, while intact fibers that were run in parallel showed normal FCCP responses (data not shown). Visual examination of the permeabilized fibers found that they were not tolerating the higher, but physiological, temperature of 37°C compared with the intact single muscle fibers. However, we did find that at temperatures equivalent to room temperature and lower, the permeabilized fibers remained viable, which is what is expected, on the basis of previous publications (20, 30). Unfortunately, the Extracellular Flux Analyzer is designed to collect the experimental data at 37°C, making the permeabilized fiber approach challenging in the single fiber model. However, using the muscle fiber bundle model as previously described is more conducive to experiments at physiological temperatures (1, 20, 26). To appreciate this, one must consider the methodological differences between the muscle fiber bundle approach and the single-muscle fiber model. Muscle fiber bundles are isolated from a section of muscle with the muscle fibers being mechanically separated to maximize surface area, while still retaining some form of attachment to each other in a bundle formation (1). Single muscle fibers are fibers that are completely dissociated from each other after isolation of the whole muscle; thus, there is no contact with previous neighboring fibers. The advantage to the single-muscle fiber approach is the ability to culture the fibers in a multiwell format for multiple days, which is not applicable to the muscle fiber bundle approach since the bundle is nonadherent. Thus, when fiber bundles are isolated and permeabilized, it is done so in ice-cold solutions until right before the mitochondrial measures are made, which allows the bundles to tolerate physiological temperatures long enough to complete the experiments However, since single muscle fibers are maintained under culture conditions and in a multiwell format, mimicking the ice-cold buffer approach prior to the measures is not possible, which limits the amount of time we can maintain the permeabilized single fibers at physiological temperatures. Unfortunately, after permeabilizing the fibers, we were unable to run the experiments fast enough to achieve reliable data. However, it should be clear that the single muscle fibers will tolerate physiological temperatures when the sarcolemma is intact and the fibers are not permeabilized. It is well documented in the literature that when single fibers are permeabilized or “skinned,” they rapidly deteriorate at physiological temperatures; however, they remain viable at temperatures that are room temperature and below (6, 33, 36). In fact, to overcome this limitation, many investigations have examined the role of temperature on permeabilized (or “skinned”) single fibers by using a temperature jump method to rapidly raise the temperature of the fiber, or by conducting their experiments in a much shorter time span than our approach (less than 30 min) (29). Finally, others have shown that permeabilized single fibers housed at higher temperatures release significant amounts of reactive oxygen species, which is associated with reduced force production and likely contributes to the deterioration (7).

Mitochondrial function deficiency in mdx fibers.

To show application of our approach for measuring mitochondrial oxygen consumption, we examined mitochondrial respiration responses in intact single fibers isolated from mdx vs. WT animals. It has been previously demonstrated that maximal mitochondrial respiration in skeletal muscle of mdx mice is approximately twofold lower compared with WT mice, which appears to be mediated by a loss in specific mitochondrial protein content (21). In the present study, we isolated intact fibers from age-matched WT and mdx mice to determine whether we could detect a mitochondrial deficiency using our approach. The fibers from mdx mice exhibited significantly lower basal (P < 0.001) and maximal (P < 0.001) mitochondrial respiration rates compared with fibers from the WT mice (Fig. 3, A and D). In addition, we detected significantly lower (P < 0.01) ECAR rates in the mdx fibers compared with the WT mice, suggesting potential defects in glycolysis in the mdx fibers (Fig. 3B). However, when we calculated the OCR/ECAR ratio, no differences were found (data not shown). When total oxygen consumption was calculated, there was a trend (P = 0.08) for a lower response to the FCCP + pyruvate in the mdx mice (Fig. 3C). We calculated the average initial oxygen consumption rates and the peak oxygen consumption rates in the WT vs. the mdx mice (Fig. 3D). The intact muscle fibers from mdx mice demonstrated reduced initial and peak oxygen consumption rates (Fig. 3D). Further, we found that the fold-increase in oxygen consumption in response to the FCCP + pyruvate stimulation was 3.9 and 2.9 in the WT and mdx, respectively. In addition, we calculated the mitochondrial spare respiratory capacity, as previously described (15, 38), in the fibers and found a 62% reduction in mdx fibers compared with the WT fibers. Spare respiratory capacity is defined as the difference between the basal and the maximal (uncoupled) OCR and was suggested to approximate the ability of mitochondria to upregulate OCR in response to an increased demand for ATP (15, 38). Thus, using the intact single fiber model, we present results that are consistent with previous findings that the mdx murine model does exhibit mitochondrial dysfunction.

DISCUSSION

Determining mitochondrial function in skeletal muscle has become a commonly employed methodology. In these experiments, we sought to determine whether mitochondrial respiration measurements could be determined in cultured intact single muscle fibers in a novel microplate format. The use of this approach enables the investigator to make measurements with minimal disruption of the myofibers. Additionally, by using appropriate culture conditions, the intact single fibers can be maintained overnight in a standard CO2 incubator for subsequent measures (4). In these experiments, intact adult muscle fibers were isolated and cultured overnight. The plates were removed from the incubator the next morning, and mitochondrial respiration was determined in a microplate format, in which 20 different wells were assayed simultaneously. Mitochondrial oxygen consumption was induced by supplying the fibers excess concentrations of substrate (i.e., pyruvate or palmitate) and simultaneously uncoupling the mitochondria through the addition of FCCP. As expected, we found that pyruvate and palmitate were equally effective at inducing oxygen consumption, but the palmitate ultimately consumed more total oxygen over the entire measurement time. To ensure the increase in OCR was representative of mitochondrial oxygen consumption, antimycin was delivered to the fibers, resulting in complete loss of the enhanced OCR. Thus, our method provides an approach to measure mitochondrial-specific respiration in enzymatically isolated single muscle fibers from an adult animal without major disruption of the membrane.

Current approaches to quantify mitochondrial respiration either rely on mechanical disruption of the whole muscle followed by differential centrifugation or chemical permeabilzation of the sarcolemma. Recent findings have suggested the isolation of mitochondria-enriched fractions from skeletal muscle can result in alterations in mitochondrial function. In an elegant study, Picard et al. (27) found that there are alterations in the stoichiometry of proteins within the electron transport chain in response to the isolation process. In addition, the isolation process further results in enhanced reactive oxygen species production in response to substrate delivery (27). The data confirm the importance of studying mitochondrial function in preparations that result in minimal disruption to the cell. Further, Picard et al. (26) confirmed this concept in skeletal muscle taken from aged animals. Specifically, they found that using standard mitochondrial isolation conditions resulted in an overexaggeration of any mitochondrial deficiency in the aged animals compared with the same measures made in permeabilized fibers, which demonstrated no outward deficiency in mitochondrial function (26). Thus, the data of Picard et al. (26) would suggest that using a permeabilized fiber bundle approach is an advantageous approach to consider when studying mitochondrial function in skeletal muscle. Here, we provide an alternative to the muscle fiber bundle approach by using intact single muscle fibers (Fig. 1). These cells are isolated from a whole muscle with all membranes intact and can be maintained in standard cell cultures for a short duration while still retaining their phenotype (~48 h). The cells can be cultured longer with adjustments to the culture conditions (4). By using substrates that can cross the sarcolemma, we demonstrate that successful mitochondrial measures can be made in this model without disruption of the cell membrane. A limitation of the approach is that we cannot add substrates unless they are permeable to the cell membrane; thus, it is difficult to achieve certain measures. Comparisons of state 3 capacity is one of the most commonly reported parameters for assessing mitochondrial function (20); however, in the intact fiber approach, it is not possible to determine a true state 3 respiratory capacity compared with using isolated mitochondria or permeabilized fiber bundles, due to an inability to regulate ADP concentrations. We use FCCP-stimulated respiration as a means to induce a maximal respiratory response, which allows us to determine the spare respiratory capacity, defined as the difference between maximal OCR and the basal OCR (25). Thus, our approach assesses the ability of the cell to function as a whole, whereas other approaches are directly working with the electron transport machinery. As with any approach, there are inherent advantages and disadvantages to each approach.

In these experiments, we also employed a novel microplate approach. To our knowledge, this is the first demonstration using this technology to measure mitochondrial respiration in muscle fibers taken from an adult animal. Albeit, others have examined single fiber respiration measures in individual fibers in an isolated bath system (8, 35). An advantage of this system is the ability on an individual plate to supply fibers isolated from the same muscle with different substrates (i.e., pyruvate or palmitate) to drive respiration. Simultaneous measurements of mitochondrial respiration and the metabolic flexibility of the mitochondria in the muscle fibers result in enhanced throughput (see Fig. 2A). Metabolic flexibility is defined as the ability of a cell to adjust fuel oxidation to fuel availability (10). The term was originally defined by Kelley and Mandarino (17) as “the capacity to switch from lipid oxidation and high rates of FA uptake to suppressed lipid oxidation and increased glucose uptake and oxidation under insulin-stimulated conditions.” A critical aspect of this definition is the ability of the mitochondria within the tissue to oxidize substrates, and clearly, in our approach, we are measuring only a small aspect of the definition. Thus, by placing the cells in the same environment, but only changing the substrate, it is feasible to quantify the oxygen consumption rates' response to the different substrates (see Fig. 2C). This response can then be compared across conditions to determine whether the mitochondria within the fibers lack an inherent ability to use different substrates. Because of the complexity of mitochondria, there are numerous points within specific metabolic pathways that could contain a defect. For example, it is feasible to imagine that if there were a limitation in β-oxidation and the experimental protocol provided substrate to the muscle fibers that did not use β-oxidation (i.e., pyruvate), then the limitation would be overlooked. In fact, Koves et al. (18) have suggested that incomplete oxidation of fatty acids can contribute to the development of insulin resistance in skeletal muscle. Additionally, it has been suggested that decreases in carnitine concentrations could result in poor lipid utilization due to poor lipid entry into the mitochondria (24). By examining more than one type of substrate, an investigator gets a more complete understanding of the function of their mitochondria within their experimental paradigm. Further, by developing an approach using intact muscle fibers, an investigator can use this as a first-pass screen to determine whether substrate flux limitations into the cells contribute to their phenotype. Specifically, if an OCR response is reduced in the intact cells, but their response is normal, when the cells are treated with a membrane-permeable substrate (i.e., transport independent), it would suggest that the limitation may be occurring at the sarcolemma. For example, analyzing substrate flux across the cell membranes could be achieved using octanoic acid (C8), a medium-chain fatty acid, which can bypass transport-mediated movement into the cell (16). CD36/FAT is a well-described lipid transporter in the muscle (14). Thus, if the muscle is experiencing an inability to traffic lipid into the muscle, a potential defect could be determined using octanoic acid in parallel experiments with palmitic acid. If no defect was detected using octanoic acid, this would suggest a transport-mediated effect. However, if the defect remained, then it would suggest that β-oxidation may be the limiting factor. The technique is also amenable to quickly screen single muscle fibers with various pharmacological agents that may affect metabolic function. This allows examination of the agents' effect on adult cells as opposed to using cells that may exist in a more embryonic state. Overall, we believe we have identified a novel approach for measuring mitochondrial function in intact cultured adult muscle fibers.

As a proof of principle assay, we chose to analyze mitochondrial respiration measures from mdx mice, the murine model of Duchenne muscular dystrophy. While the loss of dystrophin is a primary deficit responsible for the initiation of the skeletal muscle dysfunction present in these mice, the pathogenic progression of the disease involves documented mitochondrial dysfunction. Experiments that used either saponin-permeabilized muscle fiber bundles or isolated mitochondria from the quadriceps muscle demonstrated impaired rates of maximal respiration compared with age-matched control mice (21). Using our approach, we confirmed this finding in that initial or basal rates for oxygen consumption were substantially reduced in the mdx mice compared with WT mice. In addition, there was a substantially lower maximal OCR in response to the pyruvate + FCCP stimulation in the mdx mice compared with the WT mice. It is likely that the differences that we detected are due to the described mitochondrial dysfunction and not plating differences. We were extremely careful to visually monitor the preplating and postplating of the cells. We saw no indications of overplating or underplating the cells in either condition (data not shown). However, we also feel that we would have been able to detect a plating effect in our experiments. Specifically, we induced peak respiration by providing the cells with a maximal dose of FCCP + pyruvate, which would provide a maximal response from the mitochondria present in that well. If our differences were due to a plating density response, we should have seen a similar percent increase from initial (baseline) to peak (FCCP) (Fig. 3D). However, when calculating the fold change from initial to peak, the mdx mice increase by 2.9-fold in response to stimulation while the WT increase by 3.9-fold, which suggests that we have detected a mitochondrial myopathy in the mdx using this approach. Further, when we calculate the spare respiratory capacity, we detected a 62% deficiency in the fibers isolated from the mdx mice compared with the WT mice. The spare respiratory capacity is the difference between the basal and the maximal (uncoupled) OCR and can be used as a surrogate readout for the ability of the mitochondria to increase oxygen consumption in response to an increased demand for ATP. Thus, if the difference we detected were entirely due to plating density, one would predict that when normalizing to the different starting points (i.e., initial OCR), all of the significant differences would be lost. Since previously published data have shown that a number of the respiratory chain components are significantly reduced in the mitochondria from the mdx compared with WT mice (9, 11, 21), we feel that our technique has successfully confirmed that mdx mice do exhibit poor functioning mitochondria compared with WT mice.

Interestingly, we also found a substantially lower ECAR response in the fibers from the mdx mice compared with the WT mice. ECAR is a measure of the rate of extracellular acidification, and it is expected that cells that have a high ECAR exhibit a glycolytic phenotype. Our data demonstrate that the initial ECAR between the fibers from the mdx and WT animals is equivalent; however, the peak stimulated ECAR response is significantly lower in the mdx compared with WT groups. This suggests that glycolysis in the mdx fibers may be compromised compared with the WT fibers. Indeed, Wehling-Henricks et al. (37) previously demonstrated that phosphofructokinase activity was significantly lower in the mdx mice compared with WT mice, an effect that was mediated by loss of neuronal nitric oxide synthase. Thus, our data provide a confirmatory and functional readout that suggests glycolysis may be impaired in fibers isolated from the mdx mice compared with the WT mice. To the best of our knowledge, there is very little evidence concerning glycolytic flux measures in mdx and WT mice and may be an area that needs to be further considered in the mdx phenotype.

A key consideration when using this approach includes the muscle fiber type. It is well documented that metabolic variation is large across and within different muscle fiber types (31). If using a muscle or model that is likely to have large variations in fiber type, it would be critical to consider performing immunohistochemistry or gel-based fiber typing.

Limitation of this approach.

To make accurate conclusions over experiments that demonstrate differences in OCR, it is critical to normalize the data to determine whether there is a potential internal mitochondrial deficiency or whether the difference is due to a reduction in the muscle fiber mitochondrial density. A number of normalization procedures are possible, including total protein, estimates of mitochondrial density through PCR-based techniques, or a cellular imaging-based technique. Albeit, in our intact fiber experiments, we have found that we were unable to extract enough protein from each well to accurately quantify total protein using a standard calorimetric protein assay (data not shown). Thus, it would be prudent to consider a more sensitive approach, such as PCR. Thus, if differences in mitochondrial DNA were detected between groups, it would indicate a reduction in mitochondrial volume per fiber; however, if no differences were found in the mitochondrial DNA, then it would suggest that an internal defect may exist within the mitochondria requiring further experiments. The approach we are describing provides a powerful means to rapidly determine whether there are differences in mitochondrial function in intact skeletal muscle fibers; however, the approach cannot accurately predict what the existing deficiency is without further follow-up experiments. Furthermore, we feel strongly that our results are not due to differences in the amount of material plated per well from each group. The XF24-3 Extracellular Flux Analyzer utilizes retractable probes that contain O2 sensors, which enter each well simultaneously, sealing off a small volume of medium to assess O2 consumption. Thus, because the probes measure a small area of the well and not the entire well, if our results were due to differences in material, we would have had to consistently underload one small area across an enormous number of measured wells. In our experiments to ensure reliability of measures, we visually assessed each well to ensure the well was not underloaded or overloaded for fiber number. We also performed a high number of replicates per condition from each muscle. For example, in the WT and mdx experiments, the OCR and ECAR measures are an average of 10 replicates per animal (n = 5 animals/group), which should ensure that any differences that we detect are actual physiological or biochemical differences and not due to material amount on the plates.

In conclusion, we believe we have developed an innovative way to assess mitochondrial respiration in skeletal muscle using a microplate format. This method enables maintenance of the intact single muscle fibers in standard cell culture conditions with minimal disruption. In addition, because the measures are made on live intact cells, the tissue can then be used in follow-up experiments, such as Western blot analyses, quantitative PCR, or imaging studies.

GRANTS

Grant funding is acknowledged from Rehabilitation R&D Research Enhancement Award Program and Biomedical R&D CDA-02 from the Veterans Affairs Research Service and the National Institutes of Health (NIH) (Grant RC2 NR011968 to C.W. Ward) (Grant R21 AR059913 to E. E. Spangenburg) and K. C. Jackson was supported by NIH Grant AG000268.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

Author contributions: R.A.S., K.C.J., C.W.W., and E.E.S. were responsible for conception and design of research; R.A.S., K.C.J., R.J.K., and C.W.W. performed the experiments; R.A.S., R.J.K., C.W.W., and E.E.S. interpreted the results of experiments; R.A.S., K.C.J., R.J.K., C.W.W., and E.E.S. edited and revised the manuscript; R.A.S., K.C.J., and E.E.S. approved the final version of the manuscript; K.C.J., R.J.K., and E.E.S. analyzed the data; K.C.J. and E.E.S. drafted the manuscript; E.E.S. prepared the figures.

ACKNOWLEDGMENTS

The authors thank Matt Hulver for insightful advice.

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