INTRODUCTION

Top Abstract Introduction Methods Results Discussion References

SEPTIC SHOCK SYNDROME represents a major cause of death in intensive care units and is usually associated with high cardiac output, severely reduced arterial pressure, and intense peripheral vasodilation (28). Bacterial lipopolysaccharide (LPS), the outer membrane of gram-negative bacteria, is known to play a central role in the pathogenesis of septic shock by activating the release of mediators and cytokines from various cells (36). In vitro and in vivo measurements of ventilatory muscle contractility and fatigue resistance have confirmed that endotoxemia or septic shock leads to a significant decline in contractility and fatigue resistance (4, 31). Many mediators such as prostaglandins, thromboxanes, reactive oxygen species, tumor necrosis factor, and lately nitric oxide (NO) have been proposed to cause muscle contractile dysfunction in septic shock.

NO, a highly reactive second messenger with numerous biological functions, is synthesized from L-arginine by a group of hemoproteins known as NO synthases (NOS). Three different genes that code for three NOS isoforms have been recognized. The neuronal (nNOS) and endothelial (ecNOS) isoforms, which were first identified in neuronal and endothelial cells, respectively, are known to have widespread tissue distribution in the brain, skeletal muscles, and endothelial cells (18). The third NOS isoform, also known as the inducible (iNOS) isoform, is expressed in numerous cell types in response to bacterial endotoxin and inflammatory cytokines such as tumor necrosis factor, interleukins, and interferon- (18).

There has been increasing interest in the past few years in the role of NO in LPS-mediated contractile dysfunction. We have documented that enhanced NO release contributes to diaphragmatic vascular dysfunction in endotoxemic dogs (14). Others have reported that in vivo and in vitro exposure to LPS or inflammatory cytokines leads to enhanced NO production and iNOS induction in muscle fibers and cultured myocytes (27, 34, 37). The functional significance of iNOS induction in muscle fibers was not addressed in these studies. Two recent reports in rats and guinea pigs suggest that muscle contractile dysfunction in septic shock coincides with iNOS induction and that inhibitors of NOS activity restore muscle contractility (5, 11). Both groups of investigators, however, have ignored the possibility that ecNOS and nNOS, which are constitutively expressed in normal muscle fibers (19, 20), could contribute to enhanced NO formation in septic muscles. Heightened NO production by these constitutive isoforms is likely to have a negative influence on muscle contractility (19). Whether constitutively expressed NOS isoforms contribute to enhanced NO production and muscle contractile dysfunction in septic shock remains unknown.

Another possible mediator of muscle contractile dysfunction in septic shock is peroxynitrite, a highly reactive compound formed by the reaction of NO with superoxide anions (2). Peroxynitrite exerts an adverse effect on protein and lipid functions as a result of oxidative modifications and nitration of tyrosine residues (2). In septic humans and animals, significant nitrotyrosine formation has been found in lung and cardiac tissue samples, suggesting that peroxynitrite formation plays a significant role in the pathogenesis of tissue injury (3, 21). The possibility that peroxynitrite forms in skeletal muscles during septic shock has not been investigated previously.

The main objectives of this study were to investigate, in a rat model of septic shock, 1) the contribution of NO to muscle contractile dysfunction, 2) which NOS isoform participates in enhanced muscle NO production, and 3) whether enhanced NO production is associated with peroxynitrite formation.

	METHODS

Top Abstract Introduction Methods Results Discussion References

Reagents. Materials for L-citrulline assay were purchased from Sigma Chemical (St. Louis, MO). L-[2,3-³H]arginine was purchased from Dupont. The Western blotting apparatus, buffers, precasted gels, membranes, and protein markers were obtained from Novex (San Diego, CA). Specific monoclonal antibodies for the three NOS isoforms were purchased from Transduction Laboratories (Lexington, KY). Polyclonal antinitrotyrosine antibody was obtained from Upstate Biotechnology (Lake Placid, NY). The enhanced chemiluminescence (ECL) detection kit was obtained from Amersham and the protein measurement kit from Bio-Rad (Hercules, CA).

Animal preparation. The procedures for the care and use of animals were approved by the Animal Care Committee of the Royal Victoria Hospital. Four groups (n = 6 in each group) of pathogen-free male Sprague-Dawley rats (250-300 g) were housed in the animal facility of the Royal Victoria Hospital and were studied 1 wk after arrival. Group 1 was injected with vehicle alone (normal saline, control group). Groups 2, 3, and 4 were injected with Escherichia coli LPS (serotype 055:B5, Sigma Chemical; 20 mg/kg ip) and killed by cervical dislocation 6, 12, and 24 h after injection, respectively. We determined in our preliminary experiments that this dose of E. coli LPS elicits a moderate degree of shock, which is associated with a significant decline in arterial pressure over a 24-h period and an ~30% mortality rate. Ventilatory and limb muscles were dissected and frozen quickly in liquid nitrogen. The tissues collected included the diaphragm, intercostal, gastrocnemius (red and white portions), and soleus muscles. For immunostaining, these muscles were sandwiched between liver slices and flash frozen in cold isopentane (20 s) and then immersed in liquid nitrogen and stored at 80°C.

L-Citrulline assay. Frozen muscle tissues from each animal were homogenized in six volumes (wt/vol) of homogenization buffer (pH 7.4) composed of 10 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid buffer, 0.1 mM EDTA, 1 mM dithiothreitol, 1 mg/ml phenylmethylsulfonyl fluoride, 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 10-ml prewarmed (37°C) tubes containing 100 µl of reaction buffer of the following composition: 50 mM KH₂PO₄, 60 mM valine, 1.5 mM NADPH, 10 mM FAD, 1.2 mM MgCl₂, 2 mM CaCl₂, 1 mg/ml bovine serum albumin, 1 µg/ml calmodulin, 10 µM tetrahydrobiopterin, and 25 µl of 120 µM stock L-[2,3-³H]arginine (150-200 cpm/pmol). The samples were incubated for 30 min at 37°C, and the reaction was terminated by the addition of 500 µl of cold (4°C) stop buffer (pH 5.5, 100 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 12 mM EDTA). To obtain free L-[³H]citrulline for the determination of enzyme activity, 2 ml of Dowex 50w resin (8% cross-linked, Na⁺ form) were added to eliminate excess L-[2,3-³H]arginine. The supernatant was removed and examined for the presence of L-[³H]citrulline by liquid scintillation counting. Enzyme activity was expressed in picomoles of L-citrulline produced per minute per milligram of total protein. Protein was measured by the Bradford technique with bovine serum albumin as standard. To differentiate between iNOS activity, which is independent of Ca²⁺ and calmodulin (18), and constitutive NOS isoform activity (Ca²⁺ and calmodulin dependent), NOS activity was also measured in the presence of 1.5 mM each 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 N^G-nitro-L-arginine methyl ester (NOS inhibitor). Ca²⁺/calmodulin-dependent NOS activity was calculated as the difference between activity measured in the presence of CaCl₂ and that measured in EDTA/EGTA buffer. Ca²⁺/calmodulin-independent NOS activity was calculated as the difference between samples assayed in the presence of EGTA/EDTA and in the presence of N^G-nitro-L-arginine methyl ester.

Immunoblotting. Crude muscle homogenate proteins (100 µg; see above) from three animals in each group were heated for 15 min at 90°C and then loaded on gradient (4-12%) sodium dodecyl sulfate-tris(hydroxymethyl)aminomethane glycine polyacrylamide gels. The proteins were transferred electrophoretically to polyvinylidene difluoride membranes, blocked with 5% nonfat dry milk, and incubated overnight at 4°C with primary monoclonal anti-iNOS, anti-ecNOS, and anti-nNOS antibodies (1:500 for each antibody). Our previous studies indicated that NOS antibodies specifically detect NOS isoforms in rat tissue samples. Lysates of cytokine-activated murine macrophages, human endothelial cells, and pituitary cells were used as positive controls for iNOS, ecNOS, and nNOS proteins, respectively (provided by Transduction Laboratories). We also evaluated the influence of LPS injection on nitrotyrosine formation (footprint of peroxynitrite) (2) by probing with a polyclonal antinitrotyrosine antibody (Upstate Biotechnology; 1 µg/ml) (2). Nitrotyrosine molecular weight standard (Upstate Biotechnology) was used as positive control. Specific proteins were detected using horseradish peroxidase-conjugated anti-mouse and anti-rabbit secondary antibodies and ECL reagents (Amersham). The blots were scanned with an imaging densitometer (model GS700, Bio-Rad; 12-bit precision and 42-µm resolution), and optical densities of the protein bands were quantified with SigmaGel software (Jandel Scientific, San Rafael, CA). Predetermined molecular weight standards (Novex) were used as markers.

Immunohistochemistry. Muscle tissues were flash frozen in cold isopentane (20 s) and then immersed in liquid nitrogen and stored at 80°C. Air-dried cryostat sections (10 µm) were rehydrated with phosphate-buffered saline (PBS, pH 7.4, 3-5 min) and blocked for 1 h with normal donkey or horse serum, then washed with PBS. For accurate detection of iNOS and nitrotyrosine expressions, we used monoclonal anti-iNOS (Transduction Laboratories; 20 µg/ml in PBS containing 1% bovine serum albumin) and polyclonal antinitrotyrosine (Upstate Biotechnology; 2 µg/ml) antibodies. Sections were incubated with primary antibodies overnight at 4°C. For iNOS expression, the sections were incubated with biotinylated goat anti-mouse secondary antibody and then treated with the avidin-biotin-peroxidase complex (Vector Laboratories, Burlingame, CA). Sites of immunoreaction were visualized by immersing the sections in a solution of diaminobenzidine and hydrogen peroxide. Counterstaining was performed with hematoxylin (Sigma Chemical). A similar protocol was used for negative control sections, except anti-iNOS antibody was replaced with mouse or rabbit IgG. For nitrotyrosine expression, we used Cy3-labeled goat anti-rabbit secondary antibody (Jackson ImmunoChemical). Sites of immunoreactivity were visualized with a Nikon fluorescence microscope equipped with a Cy3 filter.

Reverse transcription-polymerase chain reaction. Total RNA was extracted from frozen diaphragm samples following the method described by Chomczynski and Sacchi (8). Total RNA (1 µg) was reverse transcribed using random hexamers and Moloney murine leukemia virus reverse transcriptase (Life Technologies). Reverse transcription (RT)-generated cDNA encoding ecNOS, nNOS, and -actin (as an internal standard and positive controls) were amplified by polymerase chain reaction (PCR). RNA with no clear -actin band in the RT-PCR products (35 cycles) was discarded from further studies. ecNOS oligonucleotide primers (synthesized in the McGill University DNA Synthesis Facility) were 5'-TACGGAGCAGCAAATCCAC-3' (forward) and 5'-CAGGCTGCAGTCCTTTGATC-3' [reverse; 819 base pairs (bp)] (35). For nNOS, we used 5'-CACATTTGCATGCATGGGCTCGA-3' (forward) and 5'-CTCTGCAGCGGTATTCATTC-3' (reverse) primers, which produce a 1,025-bp PCR product (6). The primers for -actin were 5'-AACCCTAAGGCCAACCGTGA-3' (forward) and 5'-TCATGAGGTAGTCTGTCAGGT-3' (reverse; 240 bp)(30). The experimental conditions for all PCRs were initial denaturation at 95°C for 5 min, 35 cycles (94°C for 1 min, 50°C for 1 min, and 72°C for 1.5 min), and a final 10-min 72°C extension. Ethidium bromide-stained 2% agarose gels were used to separate PCR products, which were visualized under ultraviolet light. Optical densities of the DNA bands were scanned with a densitometer (see above) and quantified with SigmaGel software. To verify the accuracy of the amplified sequence, PCR products were cloned in PCRII cloning vector (Invitrogen, San Diego, CA) and sequenced in the McGill University DNA Sequencing Facility. To use RT-PCR semiquantitatively, we assessed the relationship between total RNA concentration per sample and the optical density of PCR products by varying RNA concentrations and fixing the number of PCR cycles at 35, as indicated previously (13). We chose to study total RNA concentrations of 50 and 20 ng for ecNOS and nNOS amplification, respectively, on the basis of the relationship between total RNA concentration obtained from the normal rat diaphragm and the optical density of ethidium bromide-stained PCR products after 35 cycles. For -actin, we used 50 ng of total RNA.

Diaphragmatic strip preparation. Diaphragms from the four groups of rats were surgically excised with ribs and central tendon attached and placed in an equilibrated (95% O₂-5% CO₂, pH 7.38) Krebs solution chilled at 4°C with the following composition (in mM): 118.0 NaCl, 4.7 KCl, 2.5 CaCl₂, 1.2 MgSO₄, 1 KH₂PO₄, 25 NaHCO₃, and 11.0 glucose. From the central tendon to the rib, a muscle strip (3-4 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, which was placed vertically in a double-jacket gut bath (Kent Scientific Instruments). A 4.0 silk thread was used to secure the central tendon to the isometric force transducer (Kent Scientific Instruments). The muscle strips were stimulated electrically at constant currents via platinum electrodes mounted in the muscle chamber and connected to a square-wave pulse stimulator (model S48, Grass Instruments). After an equilibration period of 30 min (22-25°C), the organ bath temperature was increased to 35°C and the maximal current necessary to elicit maximal force during 120-Hz stimulation frequency (600-ms duration) was identified. Muscle length was then gradually adjusted with a micrometer to the optimal value at which maximal isometric muscle force was generated in response to supramaximal stimulation (300- to 350-mA current, 120-Hz frequency). Muscle contractility was evaluated by stimulating the muscle at 10, 50, and 120 Hz while maintaining constant supramaximal current and stimulation duration (600 ms). In group 4 (24 h after LPS injection), muscle contractility was remeasured after 30 min of exposure to 1 mM S-methylisothiourea, a known powerful inhibitor of NOS activity (33). In our previous study we found that, at this concentration, exposure to S-methylisothiourea completely inhibited muscle NOS activity (15). Tetanic contractions were digitized at a frequency of 1 kHz with a personal computer and 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 muscle cross-sectional area estimated by using the value of 1.056 g/cm³ for muscle density (9). The peak force (N/cm²) was measured for each contraction within the force-frequency curve.

Data analysis. NOS activity and muscle force values are means ± SE. Differences in NOS activity between the control and LPS groups for a given muscle and between muscles were compared by two-way analysis of variance for repeated measures. Any differences detected were evaluated post hoc by the Student-Newman-Keuls test. P < 0.05 was considered significant.

	RESULTS

Top Abstract Introduction Methods Results Discussion References

Figure 1 illustrates the changes in total and Ca²⁺/calmodulin-independent NOS activities of various muscles in the four groups of rats. Total muscle NOS activity in the control group was dependent on the presence of Ca²⁺ and calmodulin and varied significantly among different muscles, with the gastrocnemius showing the highest activity and the soleus the lowest. The diaphragm exhibited a significant rise in total NOS activity 6 h after LPS injection (P < 0.05 compared with control values), whereas intercostal and gastrocnemius NOS activities increased significantly only after 12 and 24 h, respectively (P < 0.01 compared with control values; Fig. 1). These elevations in diaphragmatic, intercostal, and gastrocnemius NOS activity were due mainly to augmented Ca²⁺/calmodulin-dependent activity (Fig. 1). Ca²⁺/calmodulin-independent activity in the diaphragm, intercostal, and gastrocnemius muscles rose significantly only 12 h after LPS injection (P < 0.01 compared with control values). LPS injection had no significant effect on soleus NOS activity.

View larger version (26K): [in this window] [in a new window]   Fig. 1.   Changes in total NO synthase (NOS) activity of different muscles of control and lipopolysaccharide (LPS)-injected animals. Ca2+/calmodulin-independent NOS activity (filled columns) of diaphragm, intercostal, and gastrocnemius muscles was significantly (P < 0.01) higher in animals 12 h after LPS injection than in controls. LPS injection had no significant effect on soleus NOS activity. * P < 0.05, ** P < 0.01 vs. control animals.

Figure 2 depicts iNOS protein expression in the diaphragm (Fig. 2A) and intercostal (Fig. 2B) muscles of the four groups of animals. iNOS expression in both muscles peaked 12 h after LPS injection and disappeared completely within 24 h. In both muscles, extensive staining with anti-iNOS antibody was detected in muscle fibers 12 h after LPS injection (Fig. 3, A and B). No iNOS staining was found in control muscles or muscles obtained 6 h after LPS injection (data not shown).

View larger version (32K): [in this window] [in a new window]   Fig. 2.   A: immunoblotting of diaphragmatic proteins with anti-inducible NOS (iNOS) antibody. iNOS expression peaked 12 h after LPS injection. Mac lysate, lysates of cytokine-activated murine macrophages. B: immunoblotting of intercostal proteins with anti-iNOS antibody. Note similarity in time course of iNOS expression in intercostals and diaphragm.

View larger version (134K): [in this window] [in a new window]   View larger version (136K): [in this window] [in a new window]   View larger version (100K): [in this window] [in a new window]   Fig. 3.   Immunostaining of intercostal muscles with anti-iNOS antibody. Muscles were obtained 12 h after LPS injection. Selective iNOS staining was detected in numerous muscle fibers (A and B); no staining was found in negative control sections (C).

Changes in diaphragmatic ecNOS and nNOS protein expression in the four groups are shown in Fig. 4. Diaphragmatic ecNOS protein expression rose twofold 24 h after LPS injection (Fig. 4A). Similarly, diaphragmatic nNOS expression peaked (5-fold increase) 12 h after LPS injection and remained elevated after 24 h (Fig. 4B). These changes in constitutive NOS isoform expression were due to increased mRNA levels (Fig. 5). Optical densities of nNOS PCR product rose by >28- and 22-fold after 12 and 24 h of LPS injection, respectively (Fig. 5). Similarly, optical densities of diaphragmatic ecNOS mRNA increased to 140 and 190% of control values after 12 and 24 h of LPS injection, respectively. No significant changes in -actin expression were detected (Fig. 5).

View larger version (34K): [in this window] [in a new window]   Fig. 4.   A: representative immunoblot in which diaphragmatic homogenates were probed with antiendothelial cell NOS (ecNOS) antibody in controls and at different times after LPS injection. Peak ecNOS expression was detected 24 h after LPS injection. B: changes in neuronal NOS (nNOS) protein expression in diaphragmatic homogenates from different groups of animals. Note significant rise in nNOS expression within 12 h of LPS injection.

View larger version (34K): [in this window] [in a new window]   Fig. 5.   Results of reverse transcription-polymerase chain reaction analysis of total RNA obtained from diaphragm of control and LPS-injected rats. Note significant induction of nNOS mRNA expression 12 and 24 h after LPS injection. Induction of diaphragmatic ecNOS mRNA was also detected 12 and 24 h after LPS injection. -Actin mRNA expression remained unchanged.

Figure 6 shows the mean densitometry of various NOS isoforms in the diaphragm. iNOS expression rose by 100-fold 12 h after LPS injection and then declined to control levels by 24 h. In comparison, nNOS and ecNOS increased by more than five- and twofold, respectively, after 24 h.

View larger version (36K): [in this window] [in a new window]   Fig. 6.   Changes in protein densitometry of various enzymes in diaphragmatic samples obtained from control and LPS-injected animals. Each bar represents a mean value of 3 immunoblots. Note difference in time course of iNOS induction compared with expression of other NOS isoforms.

Immunoblotting of diaphragmatic muscle samples from control animals with antinitrotyrosine antibody revealed two faint bands equivalent to molecular masses of 50 and 42 kDa (Fig. 7A). In addition to these bands, two new prominent bands with apparent molecular masses of 196 and 86 kDa were detected in muscles of LPS-injected animals (Fig. 7A). Figure 7B illustrates the densitometric values of diaphragmatic and intercostal nitrotyrosine expression assessed by slot blotting. In both muscles a significant increase in nitrotyrosine intensity was detected only 12 h after LPS injection, with a later decline to values similar to those of the controls. Preincubation of antinitrotyrosine antibody with 10 mM authentic nitrotyrosine eliminated the positive bands mentioned above, confirming the specificity of the antinitrotyrosine antibody. Immunostaining with antinitrotyrosine antibody of diaphragmatic samples obtained from control animals showed weak positive staining close to the sarcolemma of several muscle fibers (Fig. 8A). Similar localization but more intense staining was detected in muscle fibers 12 h after LPS injection (Fig. 8B). In addition, intense nitrotyrosine staining of diaphragmatic muscle vessels was also noted after LPS injection (Fig. 8D). Replacement of antinitrotyrosine with rabbit IgG completely eliminated the staining (Fig. 8C). Similarly, sites of reactivity were eliminated when antinitrotyrosine antibody was preincubated with 10 mM authentic nitrotyrosine (not shown).

View larger version (43K): [in this window] [in a new window]   View larger version (27K): [in this window] [in a new window]   Fig. 7.   A: representative immunoblots in which diaphragmatic homogenates of control and LPS-injected rats were probed with antinitrotyrosine antibody. Note appearance of 2 major protein bands after LPS injection (arrowheads). B: mean densitometric values of antinitrotyrosine slot blotting of diaphragmatic and intercostal homogenates from various groups. ** P < 0.01 compared with control animals.

View larger version (100K): [in this window] [in a new window]   Fig. 8.   Immunostaining of control (A) and septic (12 h after LPS injection, B and D) diaphragms with antinitrotyrosine antibody. Weak positive staining (white areas) was detected close to sarcolemma of control muscle fibers (A). More intense staining of muscle fibers (B) and blood vessels (D) was detected after LPS injection. C: negative control section.

Figure 9 shows the influence of LPS injection on diaphragmatic contractility measured in vitro. Although 6 h of endotoxemia had no effect on the force-frequency relationship, diaphragmatic force generated in response to 50 Hz decreased significantly (P < 0.05 compared with control animals) 12 h after LPS injection (Fig. 9, middle). Submaximal diaphragmatic force declined further after 24 h and was associated with a reduction in maximal force (P < 0.05 compared with control values; Fig. 9, right). Incubation of diaphragmatic strips with S-methylisothiourea restored the decrease in submaximal diaphragmatic force (10- and 50-Hz stimulation) to values similar to those recorded in control animals, whereas the LPS-induced diminution of maximal diaphragmatic force was not altered by this NOS inhibitor (Fig. 10).

View larger version (40K): [in this window] [in a new window]   Fig. 9.   Changes in diaphragmatic contractility in response to 10-, 50-, and 120-Hz stimulation frequencies. * P < 0.05, ** P < 0.01 compared with control values. A significant decline in diaphragmatic force was detected at submaximal and maximal stimulation frequencies 24 h after LPS injection.

View larger version (21K): [in this window] [in a new window]   Fig. 10.   Influence of NOS inhibition on diaphragmatic contractility. Diaphragmatic strips isolated 24 h after LPS injection were stimulated at different frequencies before (cross-hatched bars) and after (filled bars) exposure to 1 mM S-methylisothiourea (SMT). ** P < 0.01 compared with control values. SMT exposure restored submaximal force with no influence on maximal diaphragmatic force generation.

	DISCUSSION

Top Abstract Introduction Methods Results Discussion References

The novel findings of this study are as follows. 1) Endotoxemia elicits a significant rise in ventilatory and limb muscle NOS activity, which is due to iNOS induction as well as nNOS and ecNOS upregulation. 2) Significant nitrotyrosine formation in the ventilatory muscles is detected 12 h after LPS injection and coincides with iNOS expression. 3) NOS inhibition completely restores the LPS-induced decline in submaximal force generation, whereas maximal muscle force remained unchanged.

iNOS expression in skeletal muscles. iNOS expression in normal skeletal muscles of various species has been investigated recently. iNOS protein expression was not detected in different skeletal muscles of pathogen-free rats (10, 12). The current study confirms the absence of iNOS protein expression in normal rat muscles. No significant iNOS expression was detected in the ventilatory and limb muscles of dogs and rabbits (15). Similarly, cultured C2C12 cells and skeletal muscles from pathogen-free C3H/HeN mouse skeletal muscles do not express iNOS protein under normal conditions (34, 37). Gath et al. (11), on the other hand, noted detectable iNOS protein expression in skeletal muscles of normal guinea pigs. These results suggest that constitutive iNOS expression in normal muscle fibers is species dependent. The functional significance of constitutive iNOS expression in guinea pig muscles remains to be assessed.

Peroxynitrite formation. It has been established that a diffusion-limited reaction of NO with superoxide anions produces peroxynitrite, which is a powerful oxidant capable of oxidizing many proteins, leading to the addition of a nitro group to the ortho position of tyrosine to form nitrotyrosine (2). Although this reaction occurs spontaneously, low-molecular-mass transition metals and superoxide dismutase are known to catalyze it (2). More recent studies have documented extensive nitrotyrosine formation in the lungs and aorta of endotoxemic animals and in patients with acute lung injury (21, 32, 38).

nNOS and ecNOS expression. Recent studies suggest that normal skeletal muscle fibers express nNOS, which is localized beneath the sarcolemma of fast-twitch fibers (19, 26). The association of nNOS with the sarcolemma is mediated through the interaction of nNOS with the dystrophin complex (7). In addition to the sarcolemma, nNOS expression is also enriched at the muscular end plate (22). Muscle fibers are reported to express the ecNOS isoform, which has a cytoplasmic distribution and is localized mainly in muscle fibers rich in succinate dehydrogenase (20).

NO and muscle contractility. Despite recent publications dealing with muscle NOS isoform expression and iNOS induction in response to LPS injection (27, 34, 37), to the best of our knowledge, only two studies have addressed the contribution of NO to LPS-induced contractile dysfunction. Boczkowski et al. (5) were the first to report that administration of NOS inhibitor 90 min after LPS infusion significantly improved twitch and tetanic diaphragmatic force to ~75% of control values. In an another study, Gath et al. (11) assessed the fatigability of guinea pig diaphragmatic strips by repeated stimulation of the phrenic nerve. These authors reported that strips isolated from endotoxemic animals fatigued faster than normal strips and that preincubation with NOS inhibitor restored muscle fatigability to normal levels. Our findings of decreased diaphragmatic contractility 24 h after LPS injection and of restored submaximal diaphragmatic force after incubation with NOS inhibitor support the notion that enhanced NO production participates in inducing contractile dysfunction in septic animals. However, unlike the results of Boczkowski et al., we noted that inhibition of muscle NOS activity by S-methylisothiourea did not restore the LPS-induced decline in maximal diaphragmatic force (Fig. 10). This observation is consistent with the hypothesis of Kobzik et al. (19) that NO influences excitation-contraction coupling at submaximal stimulation frequencies, whereas force production at maximal stimulation frequencies is insensitive to NO.