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J Appl Physiol 98: 1420-1426, 2005. First published December 10, 2004; doi:10.1152/japplphysiol.00849.2004
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NAD(P)H fluorescence imaging of mitochondrial metabolism in contracting Xenopus skeletal muscle fibers: effect of oxygen availability

Michael C. Hogan,1 Creed M. Stary,1 Robert S. Balaban,2 and Christian A. Combs3

1Department of Medicine, University of California, San Diego, La Jolla, California; and 2Laboratory of Cardiac Energetics and 3Light Microscopy Facility, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Maryland

Submitted 9 August 2004 ; accepted in final form 9 December 2004


   ABSTRACT
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 ABSTRACT
METHODS
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 DISCUSSION
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The blue autofluorescence (351 nm excitation, 450 nm emission) of single skeletal muscle fibers from Xenopus was characterized to be originating from mitochondrial NAD(P)H on the basis of morphological and functional correlations. This fluorescence signal was used to estimate the oxygen availability to isolated single Xenopus muscle fibers during work level transitions by confocal microscopy. Fibers were stimulated to generate two contractile periods that were only different in the PO2 of the solution perfusing the single fibers (PO2 of 30 or 0–2 Torr; pH = 7.2). During contractions, mean cellular NAD(P)H increased significantly from rest in the low PO2 condition with the core (inner 10%) increasing to a greater extent than the periphery (outer 10%). After the cessation of work, NAD(P)H decreased in a manner consistent with oxygen tensions sufficient to oxidize the surplus NAD(P)H. In contrast, NAD(P)H decreased significantly with work in 30 Torr PO2. However, the rate of NAD(P)H oxidation was slower and significantly increased with the cessation of work in the core of the fiber compared with the peripheral region, consistent with a remaining limitation in oxygen availability. These results suggest that the blue autofluorescence signal in Xenopus skeletal muscle fibers is from mitochondrial NAD(P)H and that the rate of NAD(P)H oxidation within the cell is influenced by extracellular PO2 even at high extracellular PO2 during the contraction cycle. These results also demonstrate that although oxygen availability influences the rate of NAD(P)H oxidation, it does not prevent NAD(P)H from being oxidized through the process of oxidative phosphorylation at the onset of contractions.

oxygen uptake; exercise; oxidative phosphorylation; mitochondria; cellular respiration

AT THE ONSET OF MUSCLE CONTRACTIONS, the rate of mitochondrial respiration is activated in a monoexponential fashion with steady state usually achieved after 45–60 s. It has been controversial as to whether the gradual onset of mitochondrial respiration at the onset of contractions is due to oxygen limitation or due to one of the other regulators of oxidative phosphorylation being rate limiting (for review, see Ref. 40). Previous studies have suggested that oxygen availability to the mitochondria does not limit muscle oxygen uptake (O2) on-kinetics during the transition from rest to contractions at submaximal O2 in isolated Xenopus muscle fibers (19). These findings at submaximal exercise intensities support the hypothesis that the limiting factor(s) for O2 on-kinetics at these work intensities is not related to insufficient oxygen available for mitochondrial respiration but likely reside within other regulatory pathways controlling skeletal muscle oxidative metabolism. However, direct support of this hypothesis from measurements of intracellular mitochondrial oxygenation state is lacking.

There are a limited number of tools to monitor oxygen tension within single cells (42). One of the approaches is to use the mitochondrial NAD(P)H fluorescence signal that permits the intracellular monitoring of the mitochondrial reduction level in a living cell. It is generally well accepted that, if oxygen is not available within the mitochondria to accept the electrons from NAD(P)H, NAD(P)H concentration will increase (4, 16, 35, 46). However, other aspects can modify the NAD(P)H level, including workload (7, 45, 46), dehydrogenase activity (26, 39), and cytosolic-mitochondria compartmentation (3, 9, 14, 15, 23, 29, 33, 44). Thus the interpretation of NAD(P)H fluorescence in intact tissues must be made with care. Even with these limitations, mitochondrial NAD(P)H fluorescence provides one of the few measurements of the distribution of mitochondrial redox state within a single cell. In addition, the use of confocal imaging on single muscle fibers also eliminates other biasing factors that complicate the interpretation of mitochondrial NAD(P)H fluorescence data such as fiber recruitment with work, fiber type differences, and inhomogeneity of blood flow and oxygen delivery that are inherent in whole animal studies.

The purpose of the present study was 1) to evaluate the use of confocal fluorescence microscopy to measure the distribution of NAD(P)H within single isolated Xenopus skeletal muscle fibers during contractile activity; 2) to examine the effect of extracellular oxygen availability on the intracellular distribution of NAD(P)H during active contractions in these muscle fibers; and 3) to test the hypothesis that oxidative phosphorylation is not oxygen limited at the onset of contractions and that the delay of oxidative phosphorylation in reaching a new steady state of ATP generation at contractile onset is a result of other intrinsic factors.


   METHODS
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 METHODS
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Experimental preparation.   The protocols for removing skeletal muscle fibers from Xenopus were approved by the Animal Use and Care Committee of the National Heart, Lung, and Blood Institute at the National Institutes of Health. Adult female Xenopus laevis were doubly pithed and decapitated. Lumbrical muscles II–IV were removed, and single living muscle fibers (n = 6) were microdissected from the muscle. Highly oxidative fibers were dissected by dark-field illumination techniques (41). Dissections and experiments were performed in Ringer solution (112 mM NaCl, 1.87 mM KCl, 0.82 mM CaCl2, 2.38 mM NaHCO3, 0.07 mM NaH2PO4, 0.1 mM EGTA) at 20°C and 7.2 pH. After dissection, platinum clips were attached to the tendons, and the fibers were mounted in a glass chamber. One tendon end was then attached to a force transducer system (Aurora Scientific, model 400A, Aurora, Ontario, Canada) for measurement of force development.

Before each experiment, the fiber length was adjusted to achieve maximal force development for a single twitch contraction (Grass S48 stimulator, Quincy, MA). Waveforms were recorded by use of an analog-to-digital converter system and a laptop computer. A Biopac Systems MP100WSW (Santa Barbara, CA) analog-to-digital converter was used to transform the analog force signal, and the digital data were collected and analyzed with AcqKnowledgeIII 3.2.6 software (Biopac Systems). Tetanic contractions were induced by end-to-end stimulation (50 impulses/s of 1-ms duration at 9 V, with a train duration of 250 ms). Low PO2 Ringer perfusate (PO2 = 0–2 Torr; PCO2 = 40 Torr) was generated by aeration with 95% N2 and 5% CO2, and high PO2 perfusate (PO2 of ~30 Torr; PCO2 = 40 Torr) was generated by aeration with 4% O2, 5% CO2, and the balance N2. The value of PO2 was chosen as representative of a mean capillary PO2 that would surround working muscle fibers in vivo. The PO2 of the Ringer solution in the chamber was monitored with a Clark-style electrode (model 733, Diamond General, Ann Arbor, MI) placed adjacent to the working fiber.

Experimental protocol.   Fibers were stimulated for two contractile periods with each period being 2 min of tetanic contractions at one contraction every 2 s separated by 60 min of rest. The two contractile periods differed only in the PO2 of the Ringer solution perfusing the single fibers. Constant perfusion was maintained during each contractile period to maintain the experimental PO2 and to reduce the possible occurrence of unstirred layers surrounding the cell. Each fiber had its rate of fatigue development measured during the two separate work bouts. Fatigue was determined as the percent fall in developed force, compared with the highest peak force within that 2-min contractile period and was reported as a relative percentage.

Confocal images of the autofluorescence of NAD(P)H [NAD(P)HF; 351 nm excitation, 450 nm emission] were collected by use of a Zeiss LSM 510 confocal microscope and a x40 (0.65 N.A.) objective. Image analysis was performed using custom-written programs in the IDL programming language (RSI, Boulder, CO). Collection of the fluorescence images was gated to the stimulation frequency so that an image was obtained every 2 s between each contraction. Images were collected from the center of the cell with 512 x 512 pixels and a 12-bit data depth and a slice thickness of 5 µM. Comparisons of NAD(P)HF distribution within each cell were made by measuring NAD(P)HF in the outer 10% of the cell (periphery) vs. the center 10% of the cell (core). Differences in the two regions were compared at 1- and 2-min time points in the contractile period. Higher resolution pixel-by-pixel comparisons of the NAD(P)H signal were not possible because of the influences of slight tissue displacement during the contraction process.

Characterization of blue fluorescence signal from Xenopus.   Figure 1 is a histological study of succinate dehydrogenase (SDH) distribution in Xenopus muscles (adapted from Ref. 41) and representative blue autofluorescence images from this study. The autofluorescence images are with a different orientation (coronal to the long axis of the fiber) because of the geometry of the apparatus. As seen in Fig. 1, the mitochondria are distributed around the sarcolemma of the type 1 fibers (larger diameter, fewer mitochondria, Xenopus fast twitch, see Fig. 1), as revealed by the SDH distribution. The blue autofluorescence had the same distribution, with the most intense fluorescence at the periphery of the type 1 fiber. In the type 3 (smaller diameter, larger volume of mitochondria, Xenopus slow twitch, see Fig. 1) fibers, the mitochondria are more evenly distributed, as seen in the SDH distribution and the blue autofluorescence from a representative type 2 fiber. This type of distribution was seen in every fiber we examined and had a striking resemblance to the published SDH distribution shown in Fig. 1. Although this pattern seems clear, because of the compromises in the optical setup to achieve the necessary working distance for this study, individual mitochondria could not be observed. In addition to the morphological correlation of the blue autofluorescence with mitochondrial distribution in the two fiber types, the blue autofluorescence also has the functional characteristics of the mitochondrial NAD(P)H pool. The addition of cyanide (~10 µM) immediately increased the blue fluorescence signal (Fig. 2), consistent with the response of mitochondrial NAD(P)H to the blockage of the cytochrome chain. Removal of oxygen, which should also inhibit oxidation of mitochondrial NADH, also resulted in similar results (Fig. 5). These data are consistent with the notion that the mitochondrial NAD(P)H is dominating the blue autofluorescence signal as seen in most mammalian systems (9, 11, 14, 23, 29) with the notable exception of the liver (3). The reasons for the dominance of the blue autofluorescence signal of NAD(P)H in muscle mitochondria compared with the cytoplasm are differences in concentration, fluorescence lifetime, and quenching or enhancement of NAD(P)H fluorescence by local proteins. The relative contribution of mitochondrial NADH or NAD(P)H is unknown, and therefore the signal could be a combination from both of these metabolites. On the basis of these data and conclusions drawn from the literature, we are assuming that the blue autofluorescence from Xenopus skeletal muscle is primarily mitochondrial NAD(P)H.



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  		Fig. 1. Mitochondrial distribution in various fiber types of Xenopus as seen by histological sections (A, B, C) and NADH autofluorescence (C, D). A, B, and C are reprinted with permission (41) and represent histological cross sections from 3 single muscle fibers stained for the mitochondrial enzyme succinate dehydrogenase: Xenopus type 1 (A), Xenopus type 2 (B), and Xenopus type 3 (C). The scale bar represents 0.1 mm. D and E are confocal images of blue autofluorescence (351 nm excitation, 450 nm emission) taken through the long axis of the muscle fiber for type 1 and 2 fibers, respectively. Details of confocal imaging methods can be found in the text.

 


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  		Fig. 2. Effect of potassium cyanide (KCN) on the blue autofluorescence from Xenopus skeletal muscle. The confocal images were taken through the center of the muscle cell along the long axis before and after the addition of KCN (1 mm). Details of confocal imaging methods can be found in the text. The graph shows the time course of the effect of KCN on this muscle fiber.

 


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  		Fig. 5. Relative changes in NAD(P)HF in Xenopus isolated single skeletal muscle fibers (n = 6) during steady-state contractions at high (30 Torr) and low (0–2 Torr) extracellular PO2. Arrows indicate the times represented in individual images in Fig. 4.

 
Statistics.   Repeated-measures analysis of variance was performed for the statistical analyses. In all analyses, the P < 0.05 level of significance was used. Results are reported as means ± SE.


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The change in relative force production in the Xenopus isolated single skeletal muscle fibers (n = 6) during the 120-s contractile period at high (30 Torr) and low (0–2 Torr) extracellular PO2 is illustrated in Fig. 3. The maximal absolute force development at the initiation of the contractile periods was not statistically different between the two conditions and is normalized to 100%. However, the fall in force over the 120-s contractile period was significantly greater (P < 0.01) in the low oxygenation condition compared with the high (61 ± 9 vs. 81 ± 9% of peak force).



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  		Fig. 3. Change in relative force production during the 120-s contractile period at high (30 Torr) and low (0–2 Torr) extracellular PO2. Inset shows the statistical summary of the data at 120 s. *P < 0.05.

 
Figure 4 illustrates a time course of confocal NAD(P)H images of the single Xenopus fibers under the high (2A) and low (2B) oxygenation conditions. The first image in both time courses is a representative control image; the following images are during and after the start of the contraction period. With high oxygen tensions the NAD(P)H fell during the contraction period, whereas the NAD(P)H increased in the low oxygen condition. These data are quantitatively summarized over the entire cell in Fig. 5. In the low PO2 muscles, there was a 6- to 8-s time delay before NAD(P)H increased. After the initial time delay, NAD(P)H rose rapidly in a monoexponential manner and reached peak fluorescence intensity at ~50 s and then gradually declined. At the cessation of work, the NAD(P)H signal dropped, consistent with an improvement in the oxygen tension in the tissue with the decreased energy demand and oxygen consumption. In the high PO2 condition, the time delay was ~2–4 s and was followed by a rapid and then slower fall in NAD(P)H. The decrease in NAD(P)H is consistent with sufficient oxygenation to result in a net oxidation of NAD(P)H with the increase in energy demand as seen in other systems (24). However, with the cessation of work, the NAD(P)H was further oxidized, suggesting that the oxygen tension was influencing the rate of NAD(P)H oxidation under these conditions. The changes in NAD(P)H were significantly different at all time points between the two oxygenation conditions. In all six cells, there was a brief upturn in the NAD(P)H at ~20–30 s.



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  		Fig. 4. Confocal images of the autofluorescence of NAD(P)H ([NAD(P)HF]) of a single skeletal muscle fiber from Xenopus at selected time points during steady-state concentrations at high (30 Torr) and low (0–2 Torr) extracellular PO2 conditions. A single control image is shown before contraction. Subsequent images are during the contractile activity. Note that in the low oxygen tension the NAD(P)H increased with contraction, whereas in the high oxygen condition the NAD(P)H generally decreased. The times of the image collections are shown in the summary plot in Fig. 5.

 
To analyze the NAD(P)H response across the cell in these relatively low-resolution studies, regions of interest making up the subsarcolemma 10% and a 10% region of the center of the cell were evaluated. These data are illustrated in Fig. 6 (low oxygen) and Fig. 7 (high oxygen). In the low-oxygen condition, the NAD(P)H rose faster (initial rates of 3.2 vs. 1.02% of initial fluorescence/s, R2 > 0.98 for first five time points) and to a greater extent in the core of the cell consistent with a lower oxygen tension generated in the core of the cell. The rapid oxidation of NAD(P)H was also observed at the cessation of work in both zones. In the high-oxygen conditions, the results were much more complex. In the periphery, NAD(P)H decreased much more rapidly (initial rates of 2.5 vs. 1.2% of initial fluorescence/s, R2 > 0.92 for first five time points after the decline in fluorescence after work begins) and reached a lower value than in the core. The core also revealed a large drop in NAD(P)H with the cessation of work, but in the periphery levels of NAD(P)H were essentially unaffected. These results are consistent with little or no oxygen limitation in the periphery of the cell under normoxic conditions, but a significant dependence appears in the core. It is also interesting to note that the upturn of NAD(P)H at ~20 s was most pronounced in the core of the high-oxygen-tension cells.



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  		Fig. 6. Relative changes in NAD(P)HF in the outer 10% vs. the center 10% of Xenopus isolated single skeletal muscle fibers (n = 6) during steady-state contractions at low (0–2 Torr) extracellular PO2. The increase in NAD(P)HF was significantly greater in the core vs. the periphery.

 


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  		Fig. 7. Relative changes in NAD(P)HF in the outer 10% vs. the center 10% of Xenopus isolated single skeletal muscle fibers (n = 6) during steady-state contractions at extracellular PO2 = 30 Torr. The NAD(P)HF at the 1-min time point (but not at 2 min) was significantly greater in the cell core vs. the periphery.

 

   DISCUSSION
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This study demonstrated that the origin of the blue fluorescence signal from single Xenopus muscle fibers was consistent with mitochondrial NAD(P)H on the basis of topology and functional data consistent with other tissue systems. With use of the blue autofluorescence signal from NAD(P)H, there are two main findings of this study. First, on the basis of the net decrease (oxidation) of the blue fluorescence signal from mitochondrial NAD(P)H during the initial contractions of these fibers in the high extracellular PO2 treatment of 30 Torr, there was adequate oxygenation to support NAD(P)H utilization during the metabolic transition from rest to an active state. Second, the effects of work and oxygenation level were found to differentially influence oxidation rates of mitochondrial NAD(P)H in a regional manner. These data show that, with increases in work, the changes in NAD(P)H were consistent with an influence of oxygen availability at the core of the fiber even at the high extracellular PO2 condition (PO2 of 30 Torr). Only the periphery of the fiber appeared to behave as a region not influenced by oxygen availability under these conditions. This is the first evidence for an oxygen availability influence on metabolism in the core of these cells, even when the calculated PO2 at the core of the cell is above that at which mitochondrial respiration is limited (34).

Studies over the last few years (20, 21, 25, 34) have speculated that intracellular PO2 influences many physiological factors in working skeletal muscle including time to fatigue, oxidative phosphorylation, and substrate levels [NAD(P)H, ADP, inorganic phosphate], and intracellular calcium dynamics even at levels well above critical partial pressure for mitochondrial function (as determined in isolated mitochondria). However, it must be noted that these studies were conducted on mammalian skeletal muscle, and it is unclear whether these observations are similar to results for Xenopus skeletal muscle. It also remains controversial as to whether oxygen limits the rate of oxidative phosphorylation during the transition from rest to work under normoxic conditions. We have demonstrated that mean intracellular PO2 falls relatively slowly at the onset of contractions, suggesting that oxygen availability within the cell is adequate during this time period (19). However, if there are cytosolic oxygen microgradients within the cell, particularly surrounding the mitochondria, then a more precise determination of mitochondrial oxygenation state is necessary to verify that oxygen is not limiting at exercise onset. To date, all studies involving measurement of PO2 for correlation with other physiological processes in muscle have been conducted using low-resolution techniques that either average oxygen tension measurements over the entire cell, average over an entire aliquot of cells, or measure oxygen tension over an area of muscle tissue. These techniques include oxygen-dependent phosphorescence quenching of palladium-porphyrin compounds (43), oxygen microelectrodes (33) myoglobin optical absorption spectra (1), myoglobin saturation as determined by cryomicrospectroscopy of frozen cell sections (17), and NMR spectroscopic determination of myoglobin saturation in whole muscle (28, 31) and single cardiac cells (37, 38).

In the present study, we have shown that that NAD(P)H autofluorescence is a reliable indicator of mitochondrial oxidative metabolism in Xenopus skeletal muscle as shown through the distribution of fluorescence signal (Fig. 1), the potassium cyanide data (Fig. 2), and from the response to hypoxic (0–2 Torr) conditions (Figs. 4–6). The high resolution of this technique allows subcellular regional estimates of oxygen availability directly at the mitochondria permitting the detection of any microgradients in the tissue. The rapid increase in NAD(P)H with work in the low-oxygen fibers, associated with the decrease in muscle performance are consistent with the notion that the NAD(P)H signal is reflecting a functional hypoxia at the mitochondria under these conditions. To support this conclusion, we found that the magnitude of change and the rate of increase of the NAD(P)H was greatest in the core compared with the periphery of the cell. This is likely due to the diffusion distances for oxygen in this myoglobin-free cell preparation resulting in an significant oxygen gradient from the core to the outside of the cell. Finally, with the cessation of work, the NAD(P)H levels fell [increase in NAD(P) concentration] consistent with the reduction in oxygen demand and return to a higher level of oxygenation. These results suggest that NAD(P)H should provide a useful marker of functional hypoxia in the Xenopus isolated single muscle fiber.

Using NAD(P)H as an assay for functional mitochondria hypoxia during work transitions with extracellular PO2 at 30 Torr, we found that the NAD(P)H levels decreased with the onset of work in both regions of the cells measured. This is consistent with levels of adequate oxygenation to support NAD(P)H utilization during the metabolic transition in the tissue from resting to a active state resulting in a net oxidation of NAD(P)H (4, 8, 24). These findings are consistent with our previous calculations, based on Fick's law of diffusion and the size of these non-myoglobin-containing fibers, that an anoxic core should not develop under the high extracellular PO2 condition (30 Torr) used in this study, even assuming maximal rates of oxygen consumption (34).

Although the oxygen partial pressure was adequate for NADH oxidation throughout the cell at the high PO2 condition, there was still clear evidence for different NAD(P)H responses to changes in workload in the outer vs. inner portions of the cells at both extracellular PO2 conditions. This is best seen with the cessation of work in the high PO2 condition in which NAD(P)H decreased (became more oxidized) in the core of the cell. This suggests that even though the net level of NAD(P)H dropped with work, its level was still influenced by the oxygenation level. That is, if oxygen was more available, the NAD(P)H level would drop further and a higher respiratory rate might be attained in the core of the fiber. However, the work-induced gradient in NAD(P)H gradient could also be influenced by regional differences in work levels (i.e., oxygen consumption) or metabolic capacities and responses. No experimental evidence for gradients in work or metabolic profiles are available for these cells; thus our working hypothesis is that this is due to oxygen diffusion alone. However, the decrease in NAD(P)H autofluorescence at the start of contractions in the high PO2 treatment suggests that oxygen is not completely limiting during this early phase of cellular respiration. Taken together, these results suggest that oxygen availability was an influencing factor in the core of these cells at these workloads under the conditions of this study.

The upturn in NAD(P)H at ~25–30 s in the normoxic conditions and, to a lesser extent, in the low PO2 condition (see Figs. 5– 7) occurred in each cell of this study. This is best seen at the 32.5-s time point in the NAD(P)H series under the higher PO2 condition shown in Fig. 5. At this time period, the steady rate of oxidation of NAD(P)H was halted and indeed reversed for a period of 5–10 s. This indicates that either NAD(P)H production or NAD(P)H utilization was altered for this short period of time before returning to a steady net decrease in NAD(P)H. In the core of the cell, this brief upturn was significantly more conspicuous, which would suggest that oxygen diffusion distances and the corresponding fall in intracellular PO2 was implicated in this phenomenon (Fig. 7). In previous studies (19, 22), it was shown that the fall in mean intracellular PO2 at the onset of contractions follows a time course similar to that seen with the decrease in NAD(P)H in this study. Also, the transient upturn of NAD(P)H occurs when mean intracellular PO2 declines to significantly low levels (as determined in other studies, see Ref. 19). We do not know what causes the transient upturn of NAD(P)H fluorescence, but we can speculate as to the most likely factors that could cause this phenomenon. These factors would include PO2 becoming limiting in the mitochondria and/or the influence of other metabolic regulators [NAD(P)H (36), Ca2+ (2, 13, 27), and inorganic phosphate (5)] on the rate of mitochondrial NAD(P)H oxidation and production. The eventual return of net NAD(P)H oxidation may be reflected by a decrease in net oxygen consumption by an adaptive decrease in work as seen in the force tracings. Clearly, the cellular mechanism for this highly reproducible transient increase in NAD(P)H is unknown and will require further investigation.

The ability to image the distribution of the NAD(P)H effects in these muscle fibers reveals that approaches that average the NAD(P)H signal over large regions may simply be averaging different zones where metabolic processes may be quite different (as shown in this study for the core and peripheral regions of the fiber). This might even be a more significant problem for in vitro multicellular mammalian systems such as the isolated papillary muscle (6) and perfused heart (4, 10, 35) where oxygen delivery might be even more restricted. Here, the oxygen surrounding the single fibers was uniform, unlike a single fiber in a whole muscle surrounded by a small number of capillaries. Under such in vivo conditions, the supply of oxygen to the mitochondria may be different than in the single fiber preparation, so it could be argued that oxygen supply to a working fiber in vivo may be more limiting. However, because the cells used in this study did not contain myoglobin, oxygen gradients within myoglobin-containing cells may be buffered. Myoglobin has been shown to be an important factor in oxygen transport in exercising muscle (12, 28, 31). Moreover, myoglobin knockout studies have shown that large compensatory changes occur in muscle systems where myoglobin has been eliminated. These changes include both functional and molecular adaptations (18, 32). Therefore, the implications of the results of this study may not be applicable to muscle systems where myoglobin is present.

In conclusion, although there was adequate oxygenation to support NAD(P)H utilization during the metabolic transition from rest to an active state, this study shows that there are regional differences involving the influence of oxygen on mitochondrial respiration in these myoglobin free muscles. In addition, there seem to be secondary modulators of the system that act in a time-dependent manner during work. Future optical studies, in particular two photon metabolic imaging (30) of these relatively large fibers, examining other metabolic effectors (such as calcium) in combination with NAD(P)H fluorescence may further define the reasons for the observed regional influence of oxygen.


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This research was supported by National Heart, Lung, and Blood Institute DIR funding and National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant AR-40155.


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. A. Combs, NHLBI Light Microscopy Facility, National Institutes of Health, 9000 Rockville Pike, Bldg. 10/Room B1D-416, Bethesda, MD 20892-1061 (E-mail: combsc{at}nhlbi.nih.gov)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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