Introduction

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NO is synthesized by three isoforms of NOS. Two of these NOS isoforms are constitutively expressed in various cells, and were first identified in brain cells (neuronal NOS; nNOS) and endothelial cells (eNOS) (1). A third isoform, inducible NOS (iNOS), has been identified in inflammatory cells and smooth muscles in response to cytokines and bacterial endotoxin (2).

nNOS has recently been identified in skeletal-muscle fibers, and occurs beneath the sarcolemma of fast-twitch fibers (3). The association of nNOS with the sarcolemma is mediated through the interaction of nNOS with the dystrophin complex (6). Disruption of the dystrophin complex in Duchenne and Becker muscular dystrophies results in downregulation of nNOS expression (7, 8), as well as displacement of nNOS from the sarcolemma to the cytoplasm (6). In addition to being enriched in the sarcolemma, nNOS expression is also enriched at the muscular endplate (9). Recent experiments indicate that skeletal-muscle nNOS is an alternatively spliced form that is expressed in brain cells (10). Muscle fibers are also reported to express eNOS, which has a cytoplasmic distribution and is localized mainly in muscle fibers rich in succinate dehydrogenase (11).

A new role for NO was recently discovered in cell differentiation and development. Inhibition of NO release in Drosophila larvae resulted in hypertrophy of organs (12). This finding, along with the transient expression of nNOS in the brain, spinal cord, and alveolar epithelial cells in mammalian embryos, is supportive of NO involvement in cell differentiation (13, 14). Lee and colleagues (15) provided first evidence of NO involvement in the fusion of chick embryo myoblasts under in vivo and in vitro conditions. Apart from these data, little is known about the regulation of various NOS isoforms during the development of mammalian skeletal muscles. The aim of this study was to investigate whether NO production and NOS-isoform expressions are regulated in fetal, neonatal, and mature mammalian muscle fibers. We also investigated the changes in the expression of argininosuccinate synthetase (AS) and argininosuccinate lyase (AL) in the developing skeletal muscle. These two enzymes are essential for the recycling of L-citrulline to L-arginine. Additionally, we compared the structure of nNOS mRNA expressed during early muscle development with that found in adult muscle fibers. We focused on the diaphragm because it is an important ventilatory muscle and because it has been well characterized during prenatal and postnatal development.

	Materials and Methods

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Reagents

Materials for the L-citrulline assay were obtained from Sigma (St. Louis, MO). L-[2,3-³H]arginine was obtained from Dupont Inc. (Mississauga, ON). Western blotting apparatus and gels were obtained from Novex Inc. (San Diego, CA). Monoclonal anti-nNOS and anti-eNOS antibodies were obtained from Transduction Laboratories (Lexington, KY). Polyclonal anti-nNOS antibody was raised against homogenous nNOS protein purified from rat cerebellum (13). The enhanced chemiluminescence (ECL) detection kit was purchased from Amersham Canada (Oakville, ON). The 100-bp DNA marker used in the study was obtained from Life Technologies Inc. (Gaithersburg, MD).

Animal Preparation

The procedures used in the study were approved by the Animal Care Committee of McGill University. Pathogen-free, time-dated, pregnant Sprague-Dawley rats at the gestational age of 18 to 21 d were obtained from Charles River Laboratories (St.-Constants, QC). Six pregnant rats were killed (gestational age: 18 d, embryonic Day 18) and diaphragms from their fetuses were quickly dissected and frozen in liquid nitrogen. Diaphragms from littermate fetuses were pooled. An additional eight pregnant rats were allowed to deliver, and their neonatal offspring of both sexes were anesthetized with pentobarbital sodium and decapitated at 1, 7, and 30 d after delivery (postnatal Days 1, 7, and 30). The fetuses' diaphragms were harvested as described earlier. Diaphragms were also obtained from adult male rats (> 115 d). For immunostaining, the diaphragms were sandwiched between liver slices and were flash frozen in cold isopentane (20 s), and then immersed in liquid nitrogen and stored at 80°C.

Diaphragm-strip Preparation

The diaphragms of adult and 7-d-old rats were surgically excised with ribs and central tendon attached, and were placed in an equilibrated (95% O₂/5% CO₂, pH 7.38) Krebs solution chilled to 4°C that had the following composition (mM): 118.0 NaCl, 4.7 KCl, 2.5 CaCl₂, 1.2 MgSO₄, 1 KH₂PO₄, 25 NaHCO₃, 0.025 d-tubocurarine, and 11.0 glucose. From the central tendon to the rib, a muscle strip (3 to 4 mm wide) was dissected free from the lateral costal portion of the diaphragm. The rib was left attached to the strip and was used to secure the diaphragm strip in the custom built Plexiglas muscle chamber. The strip was mounted in the muscle chamber and the muscle chamber was mounted vertically in a double-jacket gut bath (Kent Scientific Instruments). A No. 4.0 silk thread was used to secure the central tendon to the isometric-force transducer (Kent Scientific Instruments, Litchfield, CT).

Muscle strips were stimulated electrically at constant currents via platinum electrodes mounted in the muscle chamber, which were connected to a square-wave pulse stimulator (Model S48; Grass Instruments, Warwick, RI). After an equilibration period of 30 min (temperature: 22 to 25°C), the organ-bath temperature was increased to 35°C and the maximum current necessary to elicit maximum force at a 120-Hz stimulation frequency (600-ms duration) was identified. Muscle length was then gradually adjusted with a micrometer to the optimal value (Lo) at which maximum isometric muscle force was generated in response to supramaximal stimulation (current: 300 to 350 mA, frequency: 120-Hz). Force-frequency relationships of adult and 7-d-old diaphragms were then constructed by varying the stimulation frequency (between 10 and 120 Hz) while keeping supramaximal current and stimulation duration (600 ms) constant. A second force-frequency relationship was constructed following 30 min of exposure to 1 mM S-methylisothiourea (SMT) (a selective NOS inhibitor). Tetanic contractions were digitized at a frequency of 1 KHz with a personal computer, and the results were stored on the hard disk for later analysis. At the end of the experiment, the strip was blotted dry and weighed. Muscle length (cm) and weight (g) were measured and used to calculate the cross-sectional area. Isometric forces were normalized for estimated muscle cross-sectional area (CSA) by using the value of 1.056 g/cm³ for muscle density (16). The peak force in N/cm² was measured for each contraction within the force-frequency curve.

L-Citrulline Assay

Frozen tissues were homogenized in 6 volumes (wt/vol) of homogenization buffer (pH 7.4, 10 mM 4-(2-hydroxyethyl)-1-piperazine-N'-2-ethanesulfonic acid [HEPES] buffer, 0.1 mM ethylenediamine tetraacetic acid [EDTA], 1 mM dithiothreitol [DTT], 1 mg/ml phenylmethylsulfonyl fluoride [PMSF], 0.32 mM sucrose, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 10 µg/ml pepstatin A). The crude homogenates were centrifuged at 4°C for 15 min at 10,000 rpm. The supernatant (50 µl) was added to prewarmed (37°C) 10-ml tubes containing 100 µl of reaction buffer of the following composition: 50 mM KH₂PO₄, 60 mM valine, 1.5 mM nicotinamide adenine dinucleotide phosphate (NADPH), 10 mM flavine adenine dinucleotide, 1.2 mM MgCl₂, 2 mM CaCl₂ 1 mg/ml bovine serum albumin (BSA), 1 µg/ml calmodulin, 10 µM tetrahydrobiopterin, and 25 µl of 120 µM stock L-[2,3-³H]arginine (150 to 200 cpm/pmol). The samples were incubated for 30 min at 37°C and the reaction was terminated by the addition of cold (4°C) stop buffer (pH 5.5, 100 mM HEPES, 12 mM EDTA). To obtain free L-[³H]citrulline for the determination of enzyme activity, 2 ml of Dowex 50w resin (8% crosslinked, Na⁺ form) were added to eliminate excess L-[2,3-³H]arginine. The supernatant was assayed for L-[³H]citrulline through liquid scintillation counting. Enzyme activity was expressed in pmol of L-citrulline produced/min/mg total protein. Protein was measured with the Bradford technique, with BSA as standard (Bio-Rad Inc., Hercules, CA). NOS activity was also measured in the presence of 1.5 mM each of ethylene glycol-bis-(-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) and EDTA, which replaced CaCl₂ and calmodulin in the reaction buffer, and in the presence of 1 mM of N^G-nitro-L-arginine methyl ester (L-NAME) (an NOS inhibitor) or of 1 mM S-methylisothiourea (SMT). Ca²⁺/calmodulin-dependent NOS activity was calculated as the difference between that measured in the presence of CaCl₂ and that measured in EDTA/EGTA buffer. Ca²⁺/ calmodulin-independent NOS activity was measured in the presence of EGTA/EDTA.

Immunoblotting

Frozen tissues were homogenized in 6 volumes (wt/vol) of homogenization buffer, as mentioned previously. The homogenates were centrifuged at 4°C for 15 min at 10,000 rpm. The supernatant (100 µg total protein) was heated for 15 min at 90°C and then loaded on gradient (4 to 12%) Tris-glycine-sodium dodecylsulfate (SDS)-polyacrylamide gels. Proteins were electrophoretically transferred onto polyvinylidene difluoride membranes, blocked overnight (4°C) with 5% nonfat dry milk, and subsequently incubated with primary monoclonal anti-eNOS (1:1,000), monoclonal anti-nNOS (1:1,000), polyclonal anti-AS (1:2,000) and polyclonal anti-AL (1:500) antibodies. NOS antibodies were raised against 20.4-kD and 22.3-kD protein fragments corresponding to human eNOS and human nNOS sequences, respectively. Our extensive preliminary experiments revealed that these antibodies are selective to their corresponding NOS isoforms in rat tissues. Lysates of human endothelial cells and human pituitary were used as positive controls. Anti-AS and anti-AL antibodies were raised in rabbits against recombinant rat proteins (17). Specific proteins were detected with horseradish peroxidase-conjugated antimouse secondary antibody and ECL reagents (Amersham Canada, Oakville, ON). The blots were scanned with an imaging densitometer (Model GS700, 12-bit precision and 42-µm resolution; Bio-Rad Inc.), and optical densities of protein bands were quantified with SigmaGel software (Jandel Scientific, San Rafael, CA). Predetermined molecular-weight standards (Novex Inc.) were used as markers.

Immunohistochemistry

Air-dried cryostat sections (10 µm) were rehydrated with phosphate-buffered saline (PBS) (pH 7.4, 3 to 5 min) and then blocked for 1 h with normal donkey or horse serum. Sections were then incubated for 1 h at room temperature with primary monoclonal anti-eNOS (5 µg/ml in 1% BSA) or polyclonal anti-nNOS (1:1,000 dilution) antibody. After three washings, sections were incubated with fluorescein isothiocyanate (FITC)-labeled goat antirabbit or donkey antimouse secondary antibodies (Jackson Immunoresearch, West Grove, PA) for 1 h at room temperature. Slides were then visualized with a Nikon fluorescence microscope and photographed with a 35 mm camera (Nikon Inc.). A similar protocol was used for negative control sections except that anti-NOS antibodies were replaced by mouse or rabbit IgG.

Reverse-Transcription-Polymerase Chain Reaction

Total RNA was extracted from tissue samples following the method described by Chomczynski and Sacchi (18). One microgram of total RNA was reverse-transcribed using random hexamers and Moloney murine leukemia virus (MMLV) reverse transcriptase (Life Technologies). Two types of polymerase chain reaction (PCR) were performed. First, we used a relatively low concentration of RT-generated complementary DNA (cDNA) (5 ng) to quantify the changes in mRNA for eNOS, nNOS, and -actin (both as an internal standard and as a positive control) during muscle development. Oligonucleotide primers (synthesized in the McGill University DNA Synthesis Facility) for eNOS were forward (5'-TACGGAGCAGCAAATCCAC-3') and reverse (5'-CAGGCTGCAGTCCTTTGATC-3') primers, which yielded an 819-bp product (19). For nNOS, we used forward (5'-CACATTTGCATGCATGGGCTCGA-3') and reverse (5'-CTCTGCAGCGGTATTCATTC-3') primers that produced a 1,025-bp product. We also amplified -actin with forward (5'-AACCCTAAGGCCAACCGTGA-3') and reverse (5'-TCATGAGGTAGTCTGTCAGGT-3') primers that produced a 240-bp product (20). Our preliminary experiments indicated that 5 ng of cDNA was within the linear range of the relationship between template concentration (total RNA) and the intensity of the PCR product. Experimental conditions for all PCRs were: initial denaturation at 95°C for 5 min, followed by 35 cycles of replication (94°C for 1 min, 55°C for 1 min, and 72°C for 1.5 min). This was followed by a final 10-min extension at 72°C. Ethidium bromide-stained 2% agarose gels were used to separate PCR products, which were visualized under UV light. The optical density of DNA bands was scanned with a densitometer and quantified with SigmaGel software (Jandel Scientific). To verify the accuracy of the amplified sequence, PCR products were cloned in PCRII plasmid (Invitrogen, San Diego, CA) and sequenced in the McGill University DNA Sequencing Facility.

Second, we used five pairs of oligonucleotides (Figure 1 and Table 1) to amplify the full coding sequence of nNOS from diaphragmatic RNA extracted from 1-d-old neonates and adult rats. These oligonucleotides amplified overlapping sequences. PCR experimental conditions were the same as described previously, except that 1 µg of first-strand cDNA was used as a template. PCR products were separated in 1.2% agarose gel in Tris-acetate-EDTA (TAE) buffer and then transferred onto Hybond-N membranes (Amersham Canada). Prehybridization was performed with 0.5 M phosphate buffer (pH 7.2) containing 1 mM EDTA and 7% SDS at 65°C for 2 to 5 h. The full-length sequence of human nNOS (a generous gift from Dr. P. Marsden, University of Toronto) was labeled with [-³²P]deoxycytosine triphosphate ([-³²P]dCTP), using the T7 QuickPrime Kit (Pharmacia Biotech) for membrane hybridization (65°C overnight). Posthybridization washing with 1× standard saline citrate (SSC) and 0.5% SDS was done at room temperature for 5 min, followed by washing with the same buffer at 65°C for 2 h. Nonspecific hybridization was removed by high-stringency washing (0.1× SSC and 0.1% SDS). Autoradiography was performed with Kodak Biomax (Eastman Kodak, Rochester, NY) film at room temperature for 4 h.

View larger version (18K): [in this window] [in a new window]   Figure 1.   Structural diversity of nNOS in 1-d-old and adult diaphragms. Five sets of oligonucleotides and PCR were used to amplify the full coding sequence of nNOS cDNA. These oligonucleotides amplified overlapping sequences (top). PCR products were separated in 1.2% agarose gel and hybridized with a full-length sequence of human nNOS (a generous gift from Dr. P. Marsden, University of Toronto). A similar procedure was used with rat cerebellar samples (lane A in the lower panel). The products of F1R1, F2R2, and F5R5 primer sets were identical in diaphragmatic and cerebellar samples. However, F3R3 and F4R4 primers amplified larger fragments in neonatal and adult diaphragms than in the cerebellum. Sequencing of the products of F3R3 and F4R4 amplification processes revealed an insert of about 102 bases (coding for 34 amino acids) that was present in neonatal and adult diaphragms (see Figure 8). This insert sequence is located between exons 16 and 17 of cerebellar nNOS.

View larger version (11K): [in this window] [in a new window]   Figure 8.   Schematic model showing predicted amino acid sequence of alternatively spliced nNOS mRNA in neonatal and adult diaphragms.

Data Analysis

Results of NOS activity and diaphragmatic force are presented as means ± SEM. Differences in NOS activity and muscle force between different age groups and different stimulation frequencies were compared through two-way analysis of variance (ANOVA) for repeated measures. Any differences detected were evaluated post hoc with the Student-Neuman-Keuls procedure. A value of P < 0.05 was considered significant.

	Results

Top Abstract Introduction Materials & Methods Results Discussion References

Diaphragmatic total NOS activity averaged 42.0 ± 5.5 pM/ min/mg protein at embryonic Day 18, and increased slightly to 44.4 ± 4.5 pmol/min/mg at postnatal Day 1 (Figure 2). NOS activity then declined progressively at postnatal Day 7, Day 30, and in adult diaphragms (Figure 2). NOS activity throughout diaphragmatic development was dependent on the presence of Ca²⁺ and calmodulin, with Ca²⁺/calmodulin-independent activity being barely detectable (Figure 2). Incubation with 1 mM of L-NAME or 1 mM of SMT completely inhibited NOS activity in all age groups. Changes in nNOS and eNOS protein expressions during diaphragmatic development are shown in Figure 3. A prominent nNOS protein band (165 kD) was detected in embryonic Day 1 diaphragm, and in several samples a smaller band (150 kD) was seen. At postnatal Day 1, nNOS protein expression rose slightly (118% of embryonic Day 18), but then declined in postnatal Day-7, Day-30, and adult muscles to about 68%, 11%, and 3%, respectively, of that measured on embryonic Day 18 (Figure 3A). By comparison, eNOS protein expression declined progressively to 51%, 20%, 18%, and 13% of its embryonic Day-18 value in postnatal Day-1, Day-7, Day-30, and adult diaphragms, respectively (Figure 3B). Modifications of nNOS and eNOS protein expression during diaphragmatic development appear to have been due to alterations in mRNA levels for these isoforms as detected by semiquantitative RT-PCR (Figure 4). Both eNOS and nNOS mRNA levels rose at postnatal Day 1 as compared with embryonic Day 18, and then declined thereafter (Figure 4).

View larger version (19K): [in this window] [in a new window]   Figure 2.   Changes in diaphragmatic NOS activity during development. E-18, Day-1, Day-7, and Day-30 refer to diaphragmatic samples obtained from embryos (embryonic Day 18) and newborn rats (postnatal Days 1, 7, and 30). *P < 0.05 and **P < 0.01 compared with E-18. Note the substantial decline in NOS activity during development.

View larger version (18K): [in this window] [in a new window]   View larger version (18K): [in this window] [in a new window]   Figure 3.   (A) Regulation of diaphragmatic nNOS protein expression during development. Monocloncal anti-nNOS antibody was used to detect nNOS protein. Note the initial rise in nNOS expression from E-18 to Day-1. nNOS expression, however, declined progressively during postnatal development. (B) Expression of endothelial NOS (eNOS) protein in the developing diaphragm. Monoclonal anti-eNOS antibody was used to detect eNOS protein. Note that eNOS expression declined progressively during postnatal development.

View larger version (54K): [in this window] [in a new window]   Figure 4.   Semiquantitative RT-PCR evaluation of eNOS and nNOS mRNA expression during diaphragmatic development. -Actin mRNA was used as a control. Five nanograms of cDNA served as template under each condition. Selective primers for nNOS sequences amplified a single fragment of 1,025 bp, whereas eNOS and -actin primers amplified 819-bp and 240-bp fragments, respectively. Note the initial rise in nNOS and eNOS mRNA expression immediately after delivery, followed by a progressive decline. No significant changes in -actin expression were seen during diaphragmatic development.

Figure 5 shows the localization of nNOS protein in 7-d-old (middle) and adult (top) diaphragms. Prominent nNOS expression was evident at the extrajunctional sarcolemma (sarcolemma other than the neuromuscular junction) of both 7-d-old and adult diaphragms (Figure 5). We detected eNOS protein only in the endothelial cells of 7-d-old and adult diaphragms (not shown).

View larger version (78K): [in this window] [in a new window]   Figure 5.   Immunostaining for nNOS protein in the adult (top) and neonatal (middle) diaphragms. Sections were probed with polyclonal anti-NOS antibody. Notice that nNOS protein in the neonatal and mature diaphragms is localized at the sarcolemma. The bottom panel indicates a negative control section in which primary anti-nNOS antibody was replaced with a rabbit IgG.

The changes in protein expression of AS and AL are shown in Figure 6. Both of these enzymes were highly expressed during the embryonic and early postnatal periods. The expression of both enzymes declined progressively during postnatal development, and only very weak expression was detected in adult muscle (Figure 6).

View larger version (21K): [in this window] [in a new window]   View larger version (17K): [in this window] [in a new window]   Figure 6.   Protein expression of argininosuccinate synthetase (AS) (A) and argininosuccinate lyase (AL) (B) in the developing skeletal muscle. Notice that strong expression was present in the neonatal and early postnatal periods; the expression of these two enzymes declined progressively during late postnatal development.

Figure 7 shows force-frequency relationships of in vitro-isolated strips from 7-d-old and adult diaphragm before and after inhibition of NOS with SMT. In the adult diaphragm, incubation with SMT elicited a significant increase in the isometric force generated in response to 10-, 20-, 30-, and 50-Hz stimulation. Maximum tetanic force was not influenced by NOS inhibition. Similarly, isometric force elicited by 10-, 20-, 30-, and 50-Hz stimulation rose significantly after NOS inhibition in 7-d-old diaphragm strips (Figure 7). When the rise in force after SMT incubation was normalized as a percentage of maximum tetanic force (Po), NOS inhibition had a relatively greater effect on force generated in response to high frequencies of stimulation (30 and 50 Hz) in the adult diaphragm than on that of 7-d-old diaphragm (Figure 7).

View larger version (25K): [in this window] [in a new window]   Figure 7.   Force-frequency relationships of in vitro isolated strips of 7-d-old and adult diaphragms before (filled circles) and after inhibition of NOS isoform with S-methylisothiourea (SMT) (open circles). In both age groups, incubation with SMT elicited a significant increase in isometric force generated in response to low-frequency stimulation. The relative increase in force (percentage of maximum tetanic force, Po) after SMT incubation was greater in the adult than in the 7-d-old diaphragm (right panel). **P < 0.01 compared with pre-SMT values; ++P < 0.01 compared with 7-d-old diaphragm.

Figure 1 shows the structural diversity of nNOS mRNA obtained from postnatal Day-1 and adult diaphragms. Using five sets of overlapping primers that spanned the full open reading frame of rat brain nNOS, we found that identical nNOS mRNA was expressed in both diaphragms. Upon cloning and sequencing of RT-PCR products, we found that F1R1, F2R2, and F5R5 primers amplified sequences in adult and newborn rat diaphragm that were identical to that of rat-brain nNOS. On the other hand, F3R3 and F4R4 primers amplified sequences in the newborn and adult diaphragm that were 102-bp larger than rat-brain nNOS. Sequencing revealed that these extra bases represented an insert between exons 16 and 17, and coded for 34 amino acids (Figure 8), as described previously for rat skeletal muscles and cultured myotubes (10). These amino acids do not share significant sequence homology with other NOS isoforms or any other known proteins.

	Discussion

Top Abstract Introduction Materials & Methods Results Discussion References

The main findings of this study are: (1) skeletal muscle NOS activity was significantly higher in the fetal and early neonatal periods than in mature muscle; (2) increased NOS activity in the fetal and neonatal muscles was due to high expression of both eNOS and nNOS, which declined significantly for both isoforms in the adult muscle; (3) AS and AL are both expressed at relatively higher levels in embryonic and neonatal muscles than in mature muscles; (4) an identical, alternatively spliced nNOS isoform was expressed in the neonatal and mature skeletal muscles; (5) nNOS was expressed at the sarcolemma both in neonatal and adult muscles; and (6) NOS activity exerts a negative influence on isometric muscle force in both neonatal and mature muscles.

NOS Expression in Skeletal Muscle

Several investigators have confirmed the existence of nNOS in skeletal-muscle fibers (3, 4, 5, 21, 22). Moreover, a positive correlation between nNOS expression and the percentage of type II fibers has also been reported in rat muscles (4, 23). Existence of the eNOS isoform in skeletal muscle fibers is still debatable. Although Kobzik and coworkers (11) described significant eNOS staining in muscle fibers rich in succinate dehydrogenase, others were unable to detect eNOS in human muscle fibers (5). Recent studies also indicate that in skeletal-muscle fibers, nNOS interacts with syntrophin, which is part of the dystrophin complex (6, 24). The absence of dystrophin in Duchenne muscular dystrophy leads to more cytoplasmic rather than sarcolemmal localization of nNOS (6).

Evidence emerging over the past few years indicates regulation of NO production during development. In the brain, blood vessels, and lungs, both nNOS and eNOS are expressed at relatively higher levels during early and late gestational periods than in adult animals (13, 25, 26). During heart development in rats, peak eNOS expression in the endocardium and in the intima of numerous small vessels was detected at embryonic Day 18 (27). Little is known about the ontogenesis of NOS isoforms in skeletal muscles. Chao and associates (8) reported that nNOS was localized at the extrajunctional sarcolemma of postnatal Day 3 and Day 7 muscles, whereas more abundant nNOS expression was seen at the junctional sarcolemma after postnatal Day 12. Our data indicate that both the eNOS and nNOS isoforms are expressed at greater levels in fetal and neonatal muscles than in mature skeletal muscle. Roles for NO in the developing muscle remain unclear. Investigators have reported that NO may influence neuromuscular transmission (9). Additionally, endogenous NO release appears to exert a negative influence on muscle force (4). Our data confirm the inhibitory effect of endogenous NO release on muscle force generation not only in mature but also in neonatal muscle. The main mechanism through which NO inhibits muscle force is likely to be increased cyclic guanosine monophosphate (cGMP) concentration and activation of cGMP-dependent kinases. These kinases are important regulators of Ca²⁺ release from the sarcoplasmic reticulum. In addition, NO is capable of attenuating Ca²⁺ influx through ryanodine receptors, inhibition of mitochondrial respiration, and regulation of creatine kinase activity (11, 28, 29).

Despite the abundance of NOS activity in the neonatal muscle, the influence of NOS inhibition on muscle force was smaller than in mature muscle. This finding, together with the localization of nNOS expression at the extrajunctional sarocelmma during development, suggests that nNOS activity in the developing muscle may mediate processes other than regulation of muscle force. One of these processes is likely to be glucose uptake (30). We also speculate that NO production in neonatal muscles may participate in myocyte differentiation. In the developing Drosophila, inhibition of NOS activity causes hypertrophy of organs, whereas ectopic expression of NOS in larvae has the opposite effect (12). These results indicate that NO acts as an antiproliferative agent during development, controlling the balance between cell proliferation and cell differentiation. An increase in NOS activity in developing lungs, blood vessels, bones, and the CNS, coinciding with the cessation of cell proliferation and the beginning of cell differentiation, tends to support this important, newly suggested role of NO (14, 25, 26, 31). Indeed, Lee and colleagues (15) showed that fusion of chick embryonic myoblasts requires the presence of NO. On the basis of these results, we propose that relatively high NO production in fetal and early neonatal muscles is necessary for myocyte differentiation, which is known to continue during the early postnatal period. The mechanisms through which NO influences muscle differentiation remain to be studied.

Our results also indicate that eNOS is regulated during muscle development in a manner similar to that reported in the developing lung and cerebral cortex (25, 26, 32). The importance of endothelial NO production in developing skeletal muscles has recently been emphasized by Pierce and coworkers, who reported a significant increase in the incidence of limb-reduction defects after prenatal inhibition of NOS in rats (33).

For its activity, NOS requires free L-arginine, which derives from a cycle in which L-citrulline is converted to L-arginine through serial catalysis by AS and AL (17). Both AS and AL are heavily expressed in liver cells, with minor expression found in various other organs (17). No information is available about the expression of these enzymes in skeletal-muscle fibers. In macrophages and smooth muscles, exposure to inflammatory cytokines elicits not only the induction of iNOS but induction of AS and AL (34, 35). Our study provides the first evidence that these two enzymes are highly expressed in embryonic and neonatal skeletal muscles. In addition, the fact that expression of AS and AL closely mimics that of constitutive NOS isoforms in developing skeletal muscles suggests that a high rate of L-citrulline turnover to L-arginine exists during skeletal-muscle development, and is designed to maintain a continuous supply of L-arginine to be used as a substrate for NOS isoforms.

Structural Diversity of NOS Isoforms

Although only three genetic NOS loci have been identified, several investigators have reported tissue-specific alternative splicing of nNOS mRNA in brain tissue (36). In addition, a small (144 kD) nNOS protein has been detected in certain areas of the brain during development (31). This small nNOS protein appears to be the product of posttranslational modifications, because only one nNOS mRNA was detected. In skeletal muscles, an alternatively spliced form of nNOS mRNA, named nNOSµ, in which a 102-bp (34 amino acid) insert is located between exons 16 and 17, has been described (10). Recently, nNOSµ transcripts have been located in the penis and lower urinary tract (37). Our study confirms the existence in neonatal and adult diaphragms of a single nNOS mRNA transcript that is identical to the nNOSµ described previously in leg muscles and differentiated myoblasts in culture (10). The functional significance of this alternatively spliced nNOS mRNA species in the regulation of NO production remains to be elucidated. Our data also indicate that eNOS cDNA sequences amplified by RT-PCR are identical in neonatal and adult diaphragms, and exhibit a high identity with the human eNOS cDNA sequence.

In summary, we found significantly increased NOS activity and constitutive NOS isoform expression during late gestational and early postnatal developmental periods in the rat diaphragm. We propose that NO plays a significant role in skeletal-muscle development.