A Fluorescence-Based Method for Measuring Nitric Oxide in Extracts of Skeletal Muscle

NITRIC OXIDE: Biology and Chemistry Vol. 5, No. 5, pp. 475– 481 (2001) doi:10.1006/niox.2001.0374, available online at http://www.idealibrary.com on

A Fluorescence-Based Method for Measuring Nitric Oxide in Extracts of Skeletal Muscle Hazel Sutherland,* Roba Khundkar,* Olga Zolle,* ,1 Anne McArdle,† Alec W. M. Simpson,* Jonathan C. Jarvis,* and Stanley Salmons* ,2 *Department of Human Anatomy and Cell Biology and †Department of Medicine, University of Liverpool, Liverpool L69 3GE, United Kingdom

Received November 15, 2000, and in revised form March 27, 2001; published online July 20, 2001

We describe here a fluorescence assay for nitric oxide synthase activity in skeletal muscle based on a new indicator, 4,5-diaminofluorescein (DAF-2). The rapid and irreversible binding of DAF-2 to oxidized NO allows real-time measurement of NO production. The method is safer and more convenient than the usual citrulline radioassay and can be used with crude muscle extracts. Rabbit fast tibialis anterior (TA) muscle had a nitric oxide synthase (NOS) activity of 44.3 ⴞ 3.5 pmol/min/mg muscle. Addition of NOS blocker N G-allyl-L-arginine reduced this activity by 43%. Slow soleus muscle displayed NOS activity of 7.3 ⴞ 2.5 pmol/min/mg muscle, 16% that of the TA muscle. Continuous stimulation of TA muscle at 10 Hz for 3 weeks reduced NOS activity by 47% to an intermediate value consistent with the associated conversion of the muscle phenotype from fast to slow. © 2001 Academic Press

Key Words: DAF-2; skeletal muscle; nitric oxide (NO); nitric oxide synthase (NOS); L-NMMA; N G-allyl-L-arginine; transformation.

1

Present address: Department of Anatomy and Developmental Biology, University College London, London WC1E 6BT, United Kingdom. 2 To whom correspondence should be addressed at Department of Human Anatomy and Cell Biology, University of Liverpool, New Medical School, Ashton Street, Liverpool L69 3GE, United Kingdom. Fax: ⫹151-794 5517. E-mail: s.salmons@liverpool. ac.uk.

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Nearly all cells are responsive to nitric oxide (1), although the precise mechanisms through which it acts are not fully established in all cases. In skeletal muscle the roles of nitric oxide (NO) 3 are believed to include control of vasodilatation (2, 3), modulation of metabolism (4, 5), and regulation of contractile function (6 –9). Two constitutive isoforms of NO synthase (NOS) are expressed in skeletal muscle fibers. The neuronal isoform (nNOS) has been demonstrated in both the cytoplasm and the sarcolemma, to an extent that varies with the species (6, 10, 11). Within the sarcolemma, affinity chromatography shows it to be associated with dystrophin (12). The endothelial isoform (eNOS) has been found to be closely associated with vascular endothelium and the mitochondria (5, 11). Under conditions of stress, skeletal muscle expresses the inducible isoform of NOS (iNOS) within capillaries, macrophages, and myocytes (13). However, much more remains to be 3

Abbreviations used: DAF-2, 4,5-diaminofluorescein; NO, nitric oxide; NOS, nitric oxide synthase; TA, tibialis anterior; G L-NMMA, N -monomethyl-L-arginine monoacetate; nNOS, neural isoform of nitric oxide synthase; eNOS, endothelial isoform of nitric oxide synthase; iNOS, inducible isoform of nitric oxide synthase; DAF-2 DA, 4,5-diaminofluorescein diacetate; EDTA, ethylenediaminetetraacetic acid; PMSF, phenylmethylsulfonyl fluoride; DDT, dithiothreitol; FMN, riboflavin monophosphate; BH 4, (6R)-5,6,7,8-tetrahydrobiopterin; FAD, flavin-adenine dinucleotide; SNAP, S-nitroso-N-acetylpenicillamine; PTI, Photon Technology International.

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learned about the roles of NO within muscle and the contribution to these roles of the various NOS isoforms. Progress in this area would be facilitated by a convenient method for analyzing the activity of the three isoforms of NOS in skeletal muscle. NOS is conventionally measured in tissue by the citrulline assay (14). However, this assay requires the handling of radioactive material and does not allow the production of NO to be followed in real time. We therefore investigated the potential for a fluorescence-based method for measuring directly the amounts of NO produced in skeletal muscle. The recently developed fluorescent indicator 4,5diaminofluorescein (DAF-2) and the membranepermeable DAF-2 diacetate (DAF-2 DA) bind rapidly and irreversibly with oxidized NO under neutral conditions to form a green-fluorescent triazole compound, DAF-2T, that can be measured spectrofluorometrically (15, 16). DAF-2 DA has been used previously as the basis of intracellular determinations of NOS activity (17–20) and, more recently, to measure real-time production of NO in bovine endothelial cells subjected to shear stress (21). Here we show that DAF-2 can be used with crude extracts of skeletal muscle to determine realtime NO production. The method was evaluated on muscle samples that could be expected to show differences in NOS activity. In small laboratory animals the level of nNOS activity is high in fibers of the fast-twitch type and low in slow-twitch fibers (6, 10). We therefore included examples of rabbit skeletal muscles in which one or the other fiber type was predominant: the tibialis anterior (TA) muscle for fast fibers and the soleus muscle for slow fibers. We also included rabbit TA muscles that had been stimulated electrically at a constant frequency of 10 Hz. The adaptive response of a fast muscle to an imposed lowfrequency pattern of activity is well documented and results in a complete phenotypic conversion from the fast to the slow type of muscle. The transformation takes place progressively over a period of weeks and involves all of the subsystems of the muscle (22–24). The variety of transitions undergone during this time provides an opportunity to examine the correlates of differences in NOS activity between fiber types and the possible role of NO in the process of

transformation. For the purposes of the present study we examined muscles that had been stimulated for 3 weeks, when the muscle was undergoing the transition from fast to slow myosin isoforms (25). In addition we examined an animal model of muscular dystrophy, the mdx mouse, a mutant in which dystrophin is not expressed and in which, therefore, nNOS is unable to associate with the sarcolemma (12). We examined 21-day-old mdx mice, whose limb muscles display maximum fiber necrosis, and 28day-old mdx mice, in which a degree of regeneration is observed (26). EXPERIMENTAL PROCEDURES

Experimental Material The experimental material consisted of rabbit TA muscle (fast-twitch); rabbit soleus muscle (slowtwitch), rabbit TA that had been subjected to continuous electrical stimulation at 10 Hz for 3 weeks, gastrocnemius muscles from 21- and 28-day-old mdx mice, and gastrocnemius muscles from control C57 mice that were age-matched to the mdx mice. Preparation of Muscle Samples Each muscle was homogenized on ice in the isolation buffer (100 mg/ml) used by Reiser et al. (27). The buffer consisted of 50 mM Tris–HCl, pH 7.4, 1 mM ethylenediaminetetraacetic acid (EDTA), and protease inhibitors (0.5 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM iodoacetamide, 1 ␮M leupeptin, 1 ␮M pepstatin A). Assay The spectrofluorometric assay was based on realtime fluorescence from the indicator DAF-2 (15, 16). A cuvette containing 1 ml of reaction mixture was monitored in a Photon Technology International (PTI) Deltascan dual excitation photometer. Muscle extracts were excited at 510 nm and fluorescence was detected at 530 nm. Both the excitation and the emission bandwidths were 5 nm. Fluorescence counts were collected at a rate of two readings per second and the data were smoothed. The gradient of

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FLUORESCENCE ASSAY FOR NO

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FIG. 1. Excitation wavelength scans (from 300 to 510 nm) for the isolation buffer and the assay buffer in the presence and absence of DAF and the NO donor SNAP. The compositions of the buffers were as described in the text. Fluorescence was collected at 530 nm with a PTI Deltascan photometer at 37°C. Bandwidths were set at 5 nm, fluorescence was recorded with scan steps of 1 nm, and the data were smoothed using PTI software.

the graph of fluorescence counts against time was then calculated with Microsoft Excel software. The assay buffer was that used by Reiser et al. (27) with some modifications. The buffer consisted of 55 mM N-2-hydroxyethylpiperazine-N⬘-2-ethane-sulfonic acid (Hepes); 20 mM Tris–HCl, pH 7.4, 2 mM L-arginine, 0.8 mM dithiothreitol (DTT), 0.4 mM EDTA, 0.8 ␮M NADPH, 1 mM MgCl2 (14), 1 mM CaCl 2 , 0.5 ␮ M calmodulin, 0.8 ␮ M riboflavin monophosphate (FMN), 6 ␮ M (6R)-5,6,7,8tetrahydrobiopterin (BH 4 ), 0.8 ␮M flavin-adenine dinucleotide (FAD), and protease inhibitors (0.2 mM PMSF, 0.4 mM iodoacetamide, 0.4 ␮M leupeptin, and 0.4 ␮M pepstatin A). To this buffer was added 100 ␮l of the muscle homogenate (final concentration, 10 mg/ml) and DAF-2 to a final concentration of 12 ␮M. DAF-2 was stored at 4°C in the presence of nitrogen to prevent oxidation of the dye. Muscles were assayed under a variety of conditions: with or without NOS blockers N Gmonomethyl-L-arginine monoacetate (L-NMMA, 100 ␮M) and N G-Allyl-L-arginine (200 ␮M), in the absence of CaCl 2, and with or without L-arginine. For purposes of evaluation and calibration, the NO donor S-nitroso-N-acetylpenicillamine (SNAP) was added to the reaction mixture as a source of NO.

Statistical Analysis Statistical comparisons were performed with Student’s unpaired t test (Instat, GraphPad Software, CA). Reagents DAF-2, L-NMMA, N G-allyl-L-arginine, and SNAP were purchased from Calbiochem–Novabiochem (UK) Ltd. (Beeston, UK). All other chemicals were purchased from Sigma–Aldrich Company, Ltd. (Dorset, UK). RESULTS

In the absence of both DAF-2 and NO, the isolation buffer showed only a basal level of fluorescence but the assay buffer showed a high level of fluorescence (Fig. 1). The reason is that to maintain NOS activity in the muscle extracts it was necessary to incorporate cofactors and substrates such as NADPH, BH 4, FMN, and FAD in the assay buffer, and these substances themselves fluoresced. It was therefore necessary to optimize the excitation and emission wavelengths for DAF-2 to minimize interference from the buffer (Fig. 1). The interfering flu-

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FIG. 2. Representative traces of NO fluorescence detected by 12 ␮M DAF-2 over a period of 25 min. Homogenized muscle samples (10 mg/ml) in 1ml of assay buffer (specified in text) were excited at 510 nm and the fluorescence was collected at 530 nm using a PTI Deltascan photometer at 37°C. Bandwidths were set at 5 nm, fluorescence was recorded at a rate of 2 readings/s, and the data were smoothed using PTI software. (A) Production of NO from TA muscle and after the addition of NOS blockers N G-monomethyl-L-arginine monoacetate (L-NMMA, 100 ␮M) and N G-allyl-L-arginine (200 ␮M). (B) NO fluorescence from three different rabbit skeletal muscle samples: control TA (fast-twitch), soleus (slow-twitch), and TA muscle stimulated for 3 weeks at 10 Hz.

orescence occurred at excitation wavelengths from 320 nm to about 480 nm. When the NO donor SNAP was added to the assay buffer containing DAF-2 a new fluorescent peak was exhibited on excitation at 510 nm that was distinct from the signal emanating from the assay buffer alone. This peak therefore corresponded to the binding of oxidized NO by DAF-2. Hence 510 nm was selected as the excitation wavelength for this study. To avoid interference between the excitation wavelength (510 nm) and the emission wavelength, stated in the literature to be

515 nm, we chose the slightly longer emission wavelength of 530 nm. This was used in the excitation wavelength scans shown in Fig. 1 and in all of the subsequent assays. In the presence of active NOS, the NO fluorescence measured under these conditions increases with time. The gradient of this increase is a measure of the level of NOS activity. Figure 2A and Table I show that the addition of two different NOS inhibitors, L-NMMA and N G-allyl-L-arginine, to samples from control TA muscles reduced the gradients by 77

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FLUORESCENCE ASSAY FOR NO TABLE I

Gradients of NO Fluorescence for Different Rabbit and Mouse Skeletal Muscle Samples Gradient Counts/second (mean ⫾ SEM) Control TA TA ⫹ L-NMMA TA ⫹ N-allyl TA (⫺) arginine TA (⫺) CaCl 2 Control soleus Stimulated TA, 3 weeks, 10 Hz Gastrocnemius C57, 21 days mdx, 21 days C57, 28 days mdx, 28 days

0.95 ⫾ 0.13 0.22 ⫾ 0.02 0.41 ⫾ 0.08 0.26 0 0.09 ⫾ 0.03 0.35 ⫾ 0.05

(n (n (n (n (n (n (n

⫽ ⫽ ⫽ ⫽ ⫽ ⫽ ⫽

7) 2) 5) 1) 1) 7) 5)

1.11 ⫾ 0.05 0.74 ⫾ 0.19 0.81 0.48 ⫾ 0.48

(n (n (n (n

⫽ ⫽ ⫽ ⫽

3) 3) 1) 2)

and 57%, respectively. These results confirm that the observed fluorescence originated from NO produced by NOS. In addition, the removal of CaCl 2 and arginine from the assay buffer for control TA muscle reduced NOS activity by 100 and 73%, respectively (Table I). Variations between the NOS activity of different muscle samples are illustrated in Fig. 2B and in the larger body of data summarized in Table I. The gradient was larger for rabbit TA muscle (0.95 ⫾ 0.13) than for either soleus muscle (0.09 ⫾ 0.03, P ⬍ 0.0001) or TA muscle stimulated for 3 weeks at 10 Hz (0.35 ⫾ 0.05, P ⬍ 0.004). Table I further suggests that the gradient for control mouse gastrocnemius muscle (1.11 ⫾ 0.05) was greater than that for muscle from a 21-day-old mdx mouse (0.74 ⫾ 0.19), although the difference did not achieve significance at P ⬍ 0.05. A differential was also observed for the 28-day mdx muscle (Table I), although this was found on more limited data. Fluorescence was converted into absolute amounts of NO by means of the NO donor SNAP, which is known to release NO on a mole per mole basis in the absence of biological material (28). A series of known concentrations of SNAP was added to assay buffer containing DAF-2 and the peak fluorescence was measured. Fluorescence originating from the assay buffer alone was subtracted from the mean data. The relationship between the concentration of SNAP and the peak fluorescence has previously been found to fit a Boltzmann sigmoidal curve

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(O. Zolle, unpublished data). Our SNAP standards provided an acceptable fit to such a curve (␹ 2 ⫽ 0.22). Where biological material is present, NO has been found to bind to DAF with an efficiency of 9.6% (16). This factor was therefore taken into account in converting gradients such as those in Fig. 2 into rates of NO production. The results of such a conversion for the experiments involving rabbit skeletal muscle samples were as follows (mean ⫾ SEM). For control TA muscle (n ⫽ 7) the rate of NO production was 44.3 ⫾ 3.5 pmol/min/mg muscle. In the presence of the NOS inhibitor N G-allyl-L-arginine (200 ␮M), this figure declined by 43% to 25.7 ⫾ 3.7 (n ⫽ 5, P ⬍ 0.01). For control soleus muscle (n ⫽ 7), NO production was 7.3 ⫾ 2.5 pmol/min/mg; this was significantly lower than the rate for control TA muscle (P ⬍ 0.001). After stimulation at 10 Hz for 3 weeks, the rate of NO production had declined to 23.6 ⫾ 2.7 (n ⫽ 5); this, too, was significantly lower than the rate for the unstimulated control TA muscle (P ⬍ 0.01).

DISCUSSION

The assay presented here has the advantage that it does not require the safety measures associated with the acquisition, use, and disposal of radioactive materials. Careful choice of excitation and emission wavelengths enabled us to measure NO despite the presence of fluorescent substrates and cofactors in the assay buffer. Using this assay, we were able to measure NOS activity in TA muscle and to confirm that inhibition with the NOS blockers L-NMMA and N G-allyl-Larginine produced the expected lowering of NOS activity (6, 27). We went on to confirm previous observations (6, 10, 27) that the level of NOS activity in mammalian slow-twitch muscle is significantly lower than that in fast-twitch muscle. After 3 weeks of stimulation at 10 Hz, NOS activity in the rabbit TA muscle was 47% that of the control. This result disagrees with the study by Reiser and colleagues (27), who observed a doubling of NOS activity after 3 weeks of stimulation at 10 Hz, but is consistent with a phenotype intermediate between that of fast and slow muscle.

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Our primary objective was to establish and to validate the fluorescent assay, and the small number of samples analyzed in other parts of the study makes it difficult to draw firm conclusions. Some of the preliminary results are, however, worthy of comment. Removal of the substrate arginine from the assay substantially suppressed NOS activity. Since endogenous levels of arginine in the muscle would be very low, this provided further confirmation that we were measuring the binding of oxidized NO to DAF. Omission of CaCl 2 from the assay of control TA muscle appeared to abolish NOS activity entirely. Both nNOS and eNOS, but not iNOS, are known to be dependent on Ca 2⫹ for activation, so this observation suggests that iNOS activity is not present at detectable levels in the control TA muscle. The remaining NOS activity shown by the mouse muscles in which the membrane-anchoring molecule dystrophin was absent was presumably attributable to nNOS that was not membrane-bound and to other isoforms of NOS. NOS activity did not appear to increase between 21 and 28 days in the mdx muscles, suggesting that NO was not involved in regenerative phenomena. Although the ratio of activity between the control TA and soleus muscles in this study is similar to that reported in the study of Reiser et al. (27), the absolute values for NOS activity in this study are higher than those reported previously. Reiser et al. (27), using the citrulline assay, recorded values of 21 and 4 pmol/min/mg protein for control TA and soleus muscles, respectively, whereas we obtained values of 44.3 and 7.3 pmol/min/mg muscle for the same two muscle types. We would not, however, wish to place undue emphasis on the differences in absolute values because of residual uncertainties in the current method of calibration with SNAP. This does not detract seriously from the utility of the assay itself. Experimental studies of NOS activity tend to focus on differences between tissues or the response to changing conditions, and in this context it is the relative, rather than the absolute, levels of NOS activity that are of interest. In this respect the fluorescent assay we have presented here would appear to be a satisfactory replacement for the existing radioactive assay.

ACKNOWLEDGMENTS H.S. was supported by the British Heart Foundation (RG/ 97001). R.K. was supported by a Physiological Society Vacation Studentship. O.Z. is a Wellcome Prize Fellow (044516/Z/98).

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