INTRODUCTION

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THE MEASUREMENT of intracellular O₂ tension (PO₂) has proven to be technically formidable (14). In skeletal muscle, a number of methods have been used to estimate intracellular PO₂ under various conditions. These include O₂ microelectrodes (18), myoglobin saturation as determined by cryomicrospectroscopy of frozen cell sections (5), and, more recently, spectroscopic relaxation determination of myoglobin saturation in whole muscle (10). Each of these techniques has value under certain applications, but none can provide a reliable measurement of intracellular PO₂ in single skeletal muscle cells over extended periods of time under conditions of rest and increased metabolic rate.

A new method has been developed for accurate measurement of PO₂ on the basis of the O₂-dependent phosphorescence quenching of palladium-porphyrin compounds (15). As the PO₂ decreases in the environment around the porphyrin compound, the phosphorescence lifetime (after a single flash of light) lengthens in a systematic manner. This technique has been used successfully for measurements of microvascular PO₂ when the porphyrin compound was carried by blood (12, 13, 21). The O₂ dependence of phosphorescence for this probe is described by the Stern-Volmer relationship

(1)

where ₀ and  are the phosphorescence lifetimes in the absence of O₂ and at a PO₂, respectively, and k_q (the quenching constant) is a second-order rate constant that is related to the frequency of collisions between O₂ and the excited triplet state of the porphyrin and the probability of energy transfer when collisions occur. To calculate the PO₂, the quenching constant and the lifetime in the absence of O₂ must be measured. These values have been well characterized for the phosphor Pd-meso-tetra (4-carboxyphenyl) porphine when bound to albumin in solution (9).

The purpose of the present study was to develop a method for measuring intracellular PO₂ in intact, isolated single skeletal muscle fibers using the O₂-dependent phosphorescence quenching of a porphyrin compound.

	METHODS

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Isolated single skeletal muscle fibers from Xenopus laevis were dissected from the lumbrical muscle and mounted in a glass chamber that contained Ringer solution at 20°C. The chamber was placed on the stage of an inverted microscope configured for epi-illumination. The preparation was observed with a Nikon Fluor objective (×40, 0.70 numerical aperture) that was used dry. A solution of 10 mM fura 2 (for visual monitoring of the injection by using excitation light at 390 nm) and 1 mM palladium-meso-tetra (4-carboxyphenyl) porphine bound to bovine serum albumin was injected into single fibers by micropipette pressure injection.

Phosphorescence quenching of the Pd-porphyrin O₂ probe was measured through a system that consisted of a flash lamp (Oxygen Enterprises), a 425-nm-band-pass excitation filter, a 630-nm cut-on emission filter, and a photomultiplier tube for collection of the phosphorescence signal. Phosphorescent-decay curves were recorded from single cells (n = 24), in which respiration had been eliminated with 2 mM NaCN while the PO₂ of the Ringer solution that surrounded the cell was varied from 0 to 159 Torr. The PO₂ of the Ringer solution that surrounded the cell was altered by infusion into the chamber of Ringer solution with a PO₂ of 0 Torr (achieved by bubbling with N₂). The PO₂ of the Ringer solution was then increased slowly from 0 to 159 Torr by slight exposure of the glass chamber that housed the fiber to the atmosphere, and the solution was stirred vigorously. The temperature of the chamber was maintained at 20°C, and the Ringer pH was kept at 7.0. A polarographic PO₂ electrode (Diamond General) was placed next to the cell to monitor extracellular PO₂.

To calculate phosphorescence lifetimes from the intracellular O₂ probe, the phosphorescence-decay curves from a series of 10 flashes (15 Hz) were averaged, and a monoexponential function was fitted to the subsequent best-fit decay curve (analysis software from Medical Systems). Between 2,000 and 3,000 points were collected from each flash, so that the average decay curve was fit from 20,000 to 30,000 points. The phosphorescence lifetime of the intracellular O₂ probe was measured every 7 s while the PO₂ of the Ringer solution in the chamber rose from 0 to 159 Torr over a period of 20-40 min. Calibration curves were determined for individual fibers, assuming that the measured value of the extracellular PO₂ (as determined with the O₂ electrode) was equal to intracellular in these respiration poisoned fibers. The measured phosphorescence-lifetime decay in a zero-O₂ environment and the relationship between measured extracellular PO₂ (and thereby intracellular PO₂) and the corresponding phosphorescence lifetime as PO₂ increased in the cell were used to calculate the phosphorescence-quenching constant k_q for the intracellular O₂ probe (see Eq. 1).

In one injected fiber, twitch contractions were initiated at a frequency of 2/s before respiration was halted by NaCN, and intracellular PO₂ was monitored.

	RESULTS

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Typical phosphorescence-decay curves are illustrated in Fig. 1 for a single isolated skeletal muscle fiber at 20°C and extracellular pH 7.0. Figure 1, A-C, shows the phosphorescence-decay curves when the extracellular PO₂ of the Ringer solution surrounding the cell was 159, 20, and 0 Torr, respectively. The phosphorescence lifetimes for this fiber were 70, 290, and 680 µs for the three different PO₂ values, respectively (Fig. 1, A-C). Each phosphorescence-decay curve was the average of 10 flashes collected in <1 s.

View larger version (17K): [in this window] [in a new window]   Fig. 1.   Phosphorescence-decay curves (each an average of 10 flashes) from a single fiber injected with porphyrin-O2 probe when extracellular PO2 was 159 Torr (A), 20 Torr (B), and 0 Torr (C). Phosphorescence lifetimes for this fiber were 70, 290, and 680 µs, respectively, for 3 different PO2 values. ch, Channels.

The correlation coefficient for fitting a monoexponential function to a typical decay curve (as shown in Fig. 1) to calculate the phosphorescence lifetime was almost always >0.97. In addition, when the collection window of the photomultiplier was moved away from the field of the injected site, whether along the length of the muscle fiber or into the Ringer solution that was devoid of tissue, the decay curve was weak and extremely rapid (<30 µs), indicating only light scattering from the flash within the Ringer solution.

Figure 2 illustrates a typical calibration run for a single fiber, with phosphorescence lifetimes calculated as the extracellular PO₂ was varied. Nitrogenated Ringer solution was added to the chamber at time 0, and vigorous stirring was initiated. After an extracellular PO₂ of 0 Torr was achieved (phosphorescence lifetime of 690 µs), the phosphorescence lifetime was measured every 7 s as the PO₂ of the chamber (as recorded periodically with the O₂ electrode) slowly rose.

View larger version (15K): [in this window] [in a new window]   Fig. 2.   Typical phosphorescence lifetime calibration plot from an isolated single fiber. Nitrogenated Ringer solution was perfused into the chamber at time 0 until extracellular PO2 reached 0 Torr (phosphorescence lifetime of 690 µs). Relationship between extracellular PO2 and phosphorescence lifetime was then monitored while extracellular PO2 slowly rose.

The relationship between the mean phosphorescence lifetime and the measured extracellular PO₂ for 24 fibers (447 points) is illustrated in Fig. 3. Using the Stern-Volmer relationship (Eq. 1), the phosphorescence lifetime in a zero PO₂ environment was 690 µs while the quenching coefficient was calculated to be 100 Torr¹ · s¹.

View larger version (17K): [in this window] [in a new window]   Fig. 3.   Relationship between mean phosphorescence lifetime and measured extracellular PO2 for 24 fibers (447 points). Using the Stern-Volmer relationship (Eq. 1), the best-fit phosphorescence lifetime in a 0-Torr PO2 environment was 690 µs, while the quenching coefficient was calculated to be 100 Torr1 · s1. Solid line depicts this relationship.

Figure 4 shows an example of using phosphorescence quenching of the porphyrin probe for measuring intracellular PO₂. The changes in the phosphorescence lifetime (Fig. 4A) and the intracellular PO₂ (Fig. 4B; using k_q and ₀ as previously determined) in a single muscle fiber at rest and steady-state work at 2 twitch contractions/s are shown. Contractions were initiated at the time period marked by the first arrow and were stopped at the second arrow. Intracellular PO₂ fell from the level of steady-state extracellular PO₂ (~28 Torr) to levels near 3-4 Torr over the 120 s of contractions, and then the levels slowly recovered back to extracellular PO₂ levels.

View larger version (16K): [in this window] [in a new window]   Fig. 4.   Change in phosphorescence lifetime (A) and calculated intracellular PO2 (B) for a single fiber in which twitch contractions (2 Hz) were initiated (arrows at left) and terminated (arrows at right). Conversion of phosphorescence lifetime to PO2 was calculated by using phosphorescence lifetime in a 0-Torr PO2 environment of 690 µs and a quenching constant of 100 Torr1 · s1.

	DISCUSSION

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The results of the present study demonstrate a novel technique for measuring intracellular PO₂ in isolated single skeletal muscle fibers.

The porphyrin probe injected into these fibers has been well characterized (9, 15) and has been used in studies for monitoring microvascular PO₂ (11-13, 21). The phosphorescence lifetime in a zero-O₂ environment for this O₂ probe has been shown to range from ~550 to 700 µs, and the quenching coefficient range is from ~200 to 300 Torr¹ · s¹, depending on the temperature and pH. The value of the phosphorescence lifetime in a zero-O₂ environment found in the present study was very similar to values expected at a pH of 7.0, which is the pH of this resting cell (1, 16), and a temperature of 20°C (9). However, the quenching coefficient k_q in the present study was somewhat lower than that found previously (9). It is likely that this was caused by the unique nature of the porphyrin probe in an intracellular environment and the background scattering of light that occurs as the excitation light passes through the Ringer solution before it illuminates the single fiber. Figure 3 shows clearly that there is a strong fit of the Stern-Volmer equation (Eq. 1), by using the calculated values of phosphorescence quenching and lifetime in zero-O₂ environment, to the measured values of PO₂ and phosphorescence lifetime.

One advantage of this porphyrin-O₂ probe is that the PO₂ signal is related to the time-dependent features of the phosphorescence signal and not to the intensity of emission (as with fluorescent signals). This eliminates many of the disadvantages inherent in fluorescent probes related to quantifying signal changes, especially if the field of view is slightly altered by movement (as may occur with muscle contractions). Furthermore, although the fitting of a single exponential to the decay curve assumes that the O₂ throughout the sampling region is homogeneously distributed (which may not always be the case in these non-myoglobin-containing cells), the typical high R² (>0.97) for the curve fitting to 10 flashes suggests that this technique is an excellent first step for quantification of intracellular PO₂. By using a highly sensitive and fast-responding charge-coupled-device imaging system, it should be possible to distinguish more adequately the distribution of O₂ throughout the cell.

Potential sources of problems with the use of this porphyrin probe include the consumption of O₂ by the quenching mechanism and the production of reactive O₂ species by the reaction of O₂ with the porphyrin probe. However, the consumption of O₂ at the concentrations of the phosphor (~5-10 µM) that diffuse throughout the intracellular environment is extremely small and, therefore, not a potential source of error. In support of this statement, if an injected cell was repeatedly subjected to rapid phosphorescence measurements over long periods of time (while extracellular PO₂ was held constant), the intracellular PO₂ did not fall as would be expected if the porphyrin were consuming substantial quantities of O₂. Because the low intracellular concentration of porphyrin probe is bound to albumin, the production of any reactive O₂ species would be small and buffered (15). In the single fibers that were injected, no long-term damage to the cells was noted when the injection was carefully done and only enough probe was injected to adequately produce a quantifiable signal. In fact, some injected fibers were kept overnight and were able to contract forcefully the next day.

The applications of this technique for measuring intracellular PO₂ to the study of skeletal muscle metabolism and function are numerous. There remain substantial ambiguities concerning the O₂ dependence of various cellular functions and the role of intracellular oxygenation in modulating the variables thought to control cell respiration and metabolism. Although it has been demonstrated that respiration in isolated mitochondria is not limited until the PO₂ falls below ~0.5 Torr (2), the O₂ dependence of intact tissue may be substantially higher (11, 19, 20). In addition, skeletal muscle fatigue is known to involve numerous variables (sometimes involving inadequate O₂ for mitochondrial respiration) that interact to reduce force development so that ATP demand does not exceed supply (4, 17). However, there is evidence that the level of tissue oxygenation can modulate metabolic processes (thereby potentially affecting fatigue development) at intracellular O₂ concentrations that are low but are not rate limiting to mitochondrial respiration (6-8). With the technique introduced in the present study, and in conjuction with measurements of cell respiration (3) and noninvasive fluorescence measurements of intracellular metabolic events (such as Ca²⁺ changes), O₂ dependency of cell function can be examined.

These results demonstrate that intracellular PO₂ can be measured in isolated single skeletal muscle fibers by using O₂-dependent quenching of an injected phosphorescent-O₂ probe. This novel technique can be used to study the O₂ dependency of cell respiration, metabolism, and function at varied skeletal muscle work intensities in an easily manipulated and well-controlled extracellular environment.