Male wild-type mice, aged 16–17 mo (adult) or 26–33 mo (old), were obtained from the Jackson Laboratory (Bar Harbor, ME) and Dr. Holly Van Remmen at the University of Texas Health Science Center, San Antonio. Mice were deeply anesthetized with an intraperitoneal injection of tribromoethanol (Avertin, 400 mg/kg). The hind paws were removed and whole lumbrical (LMB) muscles were dissected from the medial side of digit 2 while immersed in a chilled bathing solution, composition (in mM): 137 NaCl, 11.9 NaHCO₃, 5.0 KCl, 1.8 CaCl₂, 0.5 MgCl₂, 0.4 NaH₂PO₄. The mice were then euthanized by an overdose of the anesthetic followed by a bilateral thoracotomy. All experimental procedures were approved by the University Committee for the Use and Care of Animals at the University of Michigan and were in accordance with the Guide for Care and Use of Laboratory Animals (Public Health Service, 19965, NIH Pub. No. 85–23).

Details of the apparatus and procedures for assessing the contractile properties of isolated mouse LMB muscles have been described previously (Claflin and Brooks, 2008). Briefly, freshly dissected muscles were transferred to a chamber that was perfused at a rate of 2 exchanges/min with Tyrode solution (mM): NaCl, 121; KCl, 5.0; CaCl₂, 1.8; MgCl₂, 0.5; NaH₂PO₄, 0.4; NaHCO₃, 24; glucose, 5.5; EDTA, 0.10. The temperature of the solution was held at 25°C and oxygenation and pH 7.3 were maintained by bubbling with a 95% O₂, 5% CO₂ mix. Activation was by electrical stimulation via platinum plate electrodes placed on either side of and parallel to the muscle. Muscles were mounted horizontally in the chamber with one end attached to a stationary post and the other to a force transducer (Aurora Scientific, Inc., modified Model 400A). The small size of the LMB muscle permitted visualization of sarcomere-based striations using standard brightfield microscopy. This allowed real-time monitoring of sarcomere length with a high-speed video system (model 901A, Aurora Scientific, Inc.), which was used to adjust resting sarcomere length to 2.5 μm for all experiments. The LMB muscle was then stimulated continuously for 2, 5, or 10 s at 125 pulses/s, a rate sufficient to elicit maximum isometric force. Each of the stimulus pulses was 0.2 ms in duration and exceeded the intensity required for maximum force production by approximately 25%. Force and NADH fluorescence (see below) were recorded continuously throughout the protocol.

The floor of the experimental chamber was made of polished quartz, which allowed unattenuated transmission of ultraviolet (UV) excitation light. The chamber was placed on the stage of an inverted microscope (Zeiss Axiovert 100). Fluorescence was elicited by epi-illumination from a 75 W xenon lamp and detected using a photomultiplier tube (Hamamatsu, model R1527 PMT). The wavelengths for excitation were centered at 361 nm (2 nm bandwidth), selected using a diffraction grating monochromator (Deltascan 4000, Photon Technology International). The emitted fluorescence passed through a 460 nm band-pass filter (50 nm bandwidth) before reaching the photomultiplier tube. Fluorescence responses were collected from a 0.23 mm by 0.92 mm rectangular area, centered on the muscle, with the long axis of the rectangle coinciding with the long axis of the muscle. Exposure of the muscle to UV light was minimized by controlling a light-blocking shutter to open only when fluorescence measurements were actively being acquired.

After attachment to the experimental apparatus, the LMB muscle was lengthened until just taut and subjected to 3–5 single stimulus pulses of increasing amplitude, separated by 10 s. The purpose of this sequence was to determine supramaximal stimulus intensity by monitoring peak twitch forces. Sarcomere length was then set to 2.5 μm by adjusting the length of the muscle while monitoring the output of the video sarcomere length analyzer. System background fluorescence was determined by moving the microscope stage just enough to remove the muscle from the microscopic field and was subtracted from all subsequent fluorescence measurements. A minimum of 10 min was allowed to elapse between the final twitch of the sequence and any fluorescence measurements to allow NADH to return to its resting level. Simultaneous continuous recording of muscle force and fluorescence was then initiated and 3 min of baseline recording was acquired before the tetanic stimulus was applied. Following the tetanic contraction, recording continued for an additional 17 min. Sample rate was 10/s throughout. For a subset of experiments (6 of 8 adult, 4 of 10 old), the chamber perfusion was stopped and the solution was replaced with a Tyrode solution to which sodium cyanide (5 mM), an electron transport inhibitor, had been added. Introduction of the cyanide resulted in a rapid increase in fluorescence that reached a plateau level within 8–10 min (see Figure 1B). The maximum fluorescence level was interpreted as corresponding to the maximum possible ratio of NADH to total NAD (NADH + NAD⁺), or the maximally reduced mitochondrial redox state. For experiments in which the maximum fluorescence level was not determined, it was estimated using the average age-specific ratio of maximum fluorescence to resting fluorescence established in the subset of experiments in which the cyanide incubation step was carried out. Although system background fluorescence was removed from all responses (described above), no attempt was made to determine the contributions of non-NADH fluorescence through the use of uncouplers such as carbonylcyanide-p-trifluoromethoxy phenylhydrazone (FCCP) (Eng et al., 1989; Minezaki et al., 1994; Brandes and Bers, 1996; Gandra et al., 2012). Because fluorescence results are all presented after normalization by either resting or maximum fluorescence, and any non-NADH fluorescence that is present is not accounted for, non-NADH fluorescence would have the effect of reducing all reported amplitudes. The peak values, Δ-peak values, and peak-peak values presented here should therefore be considered lower-bound estimates of the true values.

Statistical analyses were performed using JMP software (SAS Institute, Inc.). Student's t-test was used to identify differences between means. Significance was set a priori at p < 0.05.

The brief, early increase in mitochondrial [NADH] following tetanic contractions (P₁) has been observed in toad skeletal muscles (Godfraind-De Becker, 1972) and in rat soleus muscles (Wendt and Chapman, 1976). Extensor digitorum longus (EDL) muscles of rats reportedly lack the initial increase in [NADH] following a tetanic contraction (Wendt and Chapman, 1976), perhaps due to differences in fiber type composition of rat soleus (predominantly type 1) and EDL (predominantly type 2) muscles (Ariano et al., 1973). The LMB muscle of the mouse hind paw contains both type 1 and type 2 fibers, although type 1 fibers occupy only ≈10% of the cross-sectional area (Smith et al., 2013). Based on this small fraction of type 1 fibers, type 2 fiber subtypes likely also contribute to the robust P₁ response observed in the present study. P₁ is abolished by iodoacetic acid, a specific inhibitor of glyceraldehyde-3-phosphate dehydrogenase (Godfraind-De Becker, 1972; Wendt and Chapman, 1976), indicating that the initial increase in mitochondrial [NADH] is a result of increased glycolytic activity, although the mechanism has not been established. Among the possibilities are increases in mitochondrial availability of pyruvate and/or the activity of NADH shuttle systems.

In cardiac muscle, an abrupt increase in energy demand by myofibrillar and sarcoplasmic reticulum ATPases causes ADP levels to rise and mitochondrial [NADH] levels to fall in response to ADP-induced activation of the electron transport system (Brandes and Bers, 2002). Recovery of mitochondrial [NADH] follows, even in the presence of continued high energy demand, as the result of a delayed increase in the activity of tricarboxylic acid (TCA) cycle dehydrogenases driven by a gradually increasing mitochondrial [Ca²⁺] (Brandes and Bers, 2002). Computational models that include [ADP]-controlled modulation of NADH oxidation rates and time-delayed [Ca²⁺]-controlled modulation of NAD⁺ reduction rates by TCA cycle activity reproduce the oscillations associated with abrupt changes in energy demand in cardiac muscle (Cortassa et al., 2006). Our interpretations of the [NADH] oscillations in skeletal muscle are based, in part, upon the findings in cardiac muscle. Specifically, we attribute the rapid decline in mitochondrial [NADH] following P₁ to ADP-induced activation of the electron transport system, resulting in an increased rate of NADH oxidation. As mitochondrial [ADP] then begins to fall due to an increase in oxidative phosphorylation, its influence on the electron transport system declines, reducing the rate of NADH oxidation by that pathway. Simultaneous with the falling NADH oxidation rate, an increase in mitochondrial [Ca²⁺] activates mitochondrial dehydrogenases, promoting an increase in the reduction of NAD⁺ to NADH by the TCA cycle. The net result of these opposing NAD oxidation and reduction activities is P₂, where mitochondrial [NADH] reaches a nadir and begins to recover. The recovery continues to P₃, where NADH oxidation rates again exceed NAD⁺ reduction rates and another local peak occurs.

The most striking differences between the NADH responses of muscles from adult and old mice are the declines in the duration and magnitude of the recovery from the minimum level (P₂) to the next local maximum (P₃). If this recovery is driven by delayed Ca²⁺-activation of TCA cycle activity as suggested by experiments on cardiac tissue (Brandes and Bers, 2002) and by computational models (Cortassa et al., 2006), then the muted response in muscles from old mice suggests diminished TCA cycle capacity or an attenuated intramitochondrial [Ca²⁺] increase in response to the preceding tetanic contraction. In support of the latter, the myoplasmic Ca²⁺ response is reduced in skeletal muscle from old humans (Delbono et al., 1995) and mice (Gonzalez et al., 2003; Umanskaya et al., 2014). Normal tetanic myoplasmic Ca²⁺ levels can be restored in old mice by overexpression of the antioxidant catalase, implicating an increase in ROS as a factor contributing to the diminished response (Umanskaya et al., 2014). Another factor that could contribute to the weak NADH recovery is increased “uncoupling” of NADH oxidation (i.e., oxidation that does not result in ATP generation), which has also been proposed to occur with aging (Conley et al., 2000; Johannsen et al., 2012). Finally, the observation that rat soleus and EDL muscles produce different NADH responses to contractions (Wendt and Chapman, 1976) raises the possibility that changes with aging in the fiber type composition contribute to the differences in NADH responses observed in the present study. We did not analyze fiber type composition of the muscles used in this study, but no changes in fiber types are observed with aging in the fore paw LMB muscle (Russell et al., manuscript under review) and minimal age-related shifts in fiber types are reported in other hind limb muscles of mice (Phillips et al., 1993; Fry et al., 2015).

A second difference between the age groups was a significant age-associated elevation in resting NADH. The observation of higher NADH for muscles of old mice is consistent with reports that the ratio of NADH to total NAD is increased in heart, lung, liver and kidney tissue from old rats (Braidy et al., 2011) and skin from older humans (Massudi et al., 2012). The elevated resting NADH indicates that the NADH-NAD⁺ pair is more reduced in old than in adult mice suggesting a more reduced mitochondrial redox environment in old mice (Mayevsky and Rogatsky, 2007; Zorov et al., 2014), although not all mitochondrial redox couples exhibit age-related changes (Dimauro et al., 2012). A more reduced mitochondrial environment in muscles of old mice increases the probability of ROS formation during cellular respiration (Zorov et al., 2014). Consistent with this possibility, mitochondria from skeletal muscles of old mice produce significantly more hydrogen peroxide (Mansouri et al., 2006), have elevated levels of regulatory proteins associated with ROS removal (Dimauro et al., 2012), and have higher antioxidant enzyme activities (Vasilaki et al., 2006) than those from adult mice. An increase in the NADH/NAD⁺ ratio is thought to be caused in part by increased activity of the NAD⁺-consuming poly (ADP-ribose) polymerase (PARP) DNA repair proteins (Braidy et al., 2011; Massudi et al., 2012), and the relative scarcity of NAD⁺ that results could have implications for other cellular enzyme systems (Sauve et al., 2006; Stein and Imai, 2012).

Monitoring mitochondrial NADH levels by measuring fluorescence offers a non-invasive means for tracking, in real-time, the mitochondrial redox state of a cell or collection of cells (Mayevsky and Rogatsky, 2007). The NADH fluorescence can thus report instantaneously the response of the mitochondrial oxidative machinery to changes in energy demand such as those that occur frequently in skeletal muscle. In addition, because the mitochondrial NADH-NAD⁺ balance is thought to reflect the global redox status of the mitochondria under study (Mayevsky and Rogatsky, 2007; Zorov et al., 2014), the fluorescence response provides valuable insights into the myriad mitochondrial functions that are affected by the redox environment and/or the availability of NADH or NAD⁺, such as calcium release by the sarcoplasmic reticulum (Zima et al., 2004), the tendency for ROS formation (Zorov et al., 2014), and the activity of sirtuin proteins (Sauve et al., 2006; Stein and Imai, 2012). Comparing fluorescence responses of skeletal muscle from adult and old mice therefore allows insights into the effects of aging on mitochondrial function and redox homeostasis and all mitochondrial processes known to be affected by redox imbalance.

Recordings of mitochondrial NADH fluctuations yield the most information when the full potential of the dynamic range of the response is known. This is typically determined by applying an inhibitor of the mitochondrial electron transport chain activity such as cyanide to maximize [NADH], and then applying an “uncoupler” such as FCCP to minimize [NADH] (Eng et al., 1989; Minezaki et al., 1994; Brandes and Bers, 1996; Gandra et al., 2012). In this study maximum NADH levels were determined by application of cyanide, but no attempt was made to assess minimum levels. Thus, an increase in non-NADH fluorescence could be contributing to the increase in resting fluorescence in muscles from old mice reported. Future experiments in which NADH levels are minimized will definitively determine resting NADH fluorescence. In addition, motion can contribute undesirable artifacts during optical recordings in muscle tissue (Godfraind-De Becker, 1972; Brandes et al., 1992; Morgan et al., 1997), but only P₁ occurs during contractile activity making TP₁ and ΔP₁ the only measurements potentially affected by motion. Finally, NADPH is excited and fluoresces at the same wavelengths as NADH, but the contribution of NADPH fluorescence to total tissue fluorescence is reported to be small (Mayevsky and Rogatsky, 2007).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.