INTRODUCTION

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It has been well established that sepsis, endotoxemia, or bacteremia is associated with significant deterioration of ventilatory muscle contractility (for review, see Hussain [1]). Several sites are influenced by these pathologies including sarcolemmal function, excitation-contraction coupling, and depression of the contractile machinery. The exact mediators responsible for the decline in ventilatory muscle contractile dysfunction in sepsis remain under investigation. Many proinflammatory cytokines and secondary mediators have been implicated such as tumor necrosis factor, products of arachidonic acid metabolism, reactive oxygen species (ROS), and lately nitric oxide (NO) (1).

NO is synthesized from L-arginine by three distinct nitric oxide synthases (NOS). In normal muscle fibers, both the neuronal (nNOS) and the endothelial (ecNOS) isoforms are responsible for NO synthesis. The nNOS isoform is localized in close proximity to the sarcolemma and associates with the dystrophin complex. This isoform has also been identified at the junctional sarcolemma and at the nerve fibers supplying skeletal muscle fibers. By comparison, the ecNOS isoform is localized at the endothelial cells of skeletal muscle vasculature and one report identified ecNOS immunoreactivity inside the sarcoplasm of type I muscle fibers (2). Unlike the other two isoforms, the iNOS isoform is not normally present inside muscle fibers but is expressed in response to the injection of bacterial lipopolysaccharide (LPS) (3).

The role of NO in the pathogenesis of sepsis-induced ventilatory and limb muscle contractile dysfunction has received some attention over the past few years. Many investigators reported that iNOS expression is induced in the ventilatory and limb muscles in response to LPS injection in various species (4). Our group has also reported that NOS activity of the ventilatory muscles increases significantly in septic rats and that both iNOS and constitutive NOS isoforms contribute to the rise in the rate of NO synthesis (3). The question of whether enhanced NO synthesis inside muscle fibers of septic animals is deleterious or protective remains unclear. Gath and coworkers (4) and Boczkowski and coworkers (6) reported that pretreatment with systemic injection of NOS inhibitors results in partial reversal of muscle contractile dysfunction as a result of iNOS inhibition. Systemic NOS inhibition has also been shown to attenuate LPS-induced ventilatory muscle sarcolemmal injury in rats (7). However, the involvement of iNOS in LPS- induced muscle dysfunction has recently been questioned by our recent finding of worsening of diaphragmatic contractile dysfunction in mice deficient in the iNOS gene (8). Although that study rules out iNOS as a mediator of ventilatory muscle contractile dysfunction, it did not address the involvement of other NOS isoforms such as the nNOS whose activity was shown to be elevated after LPS injection in iNOS-deficient mice (8).

In this study, we assessed the role of the nNOS isoform in the pathogenesis of LPS-induced diaphragmatic contractile dysfunction, iNOS induction, and sarcolemmal injury by using mice in which the nNOS gene has been disrupted (knockout). Several groups have recently used these mice to document the role of nNOS in various pathologies such as brain ischemia. Our results indicate that the nNOS isoform may play a protective role in the prevention of LPS-induced impairment of diaphragmatic force but is not a major factor in the pathogenesis of sarcolemmal injury in septic animals.

	METHODS

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Reagents

Materials for NOS activity and Escherichia coli LPS were obtained from Sigma (St. Louis, MO). Immunoblotting apparatus and reagents were obtained from Novex Inc. (San Diego, CA). Polyclonal anti-NOS antibodies were obtained from Transduction Laboratories (Lexington, KY). ECL kit was obtained from Amersham Canada Corp. (Oakville, ON, Canada).

Animal Preparation

All experiments were approved by the Animal Care Committee of McGill University. Adult (8-12 wk old) F3 and F4 C57BL/6 nNOS^/ mice were generated as mentioned previously (9). Wild-type (WT) C57BL/6 mice were purchased from Charles River Inc. Four groups of animals were studied (n = 12 in each group). Two groups of nNOS^/ mice were injected intraperitoneally with either 25 mg/kg E. coli LPS (nNOS^/-LPS) (serotype 055:B5) or an equivalent volume of normal saline (nNOS^/-C) and killed 12 h later. Similarly, two groups (WT-LPS and WT-C) of wild-type C57BL/6 mice were injected with either E. coli LPS or normal saline and killed after 12 h. LPS and saline-injected mice were matched for age, sex, and weight. At the end of the experiment, the animals were sacrificed with an overdose of sodium pentobarbital and the diaphragm was quickly excised, cleaned of connective tissue, and was either examined for contractility or sarcolemmal injury or frozen under 80° C in liquid nitrogen and prepared for immunoblotting and NOS activity.

Muscle Sample Preparation

Frozen diaphragmatic samples were homogenized in 6 volumes (wt/ vol) of homogenization buffer (pH 7.4, 10 mM HEPES buffer, 0.1 mM ethylenediaminetetraacetic acid [EDTA], 1 mM dithiothreitol, 1 mg/ml phenylmethylsulfonyl fluoride [PMSF], 0.32 mM sucrose, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 10 µg/ml pepstatin A). The crude homogenates were centrifuged at 4° C for 15 min at 10,000 rpm and the supernatants were collected.

Immunoblotting

Crude muscle homogenate proteins (80-100 µg) (see above) were heated for 15 min at 90° C and then loaded on gradient (4-12%) sodium dodecyl sulfate-tris glycine polyacrylamide gels. The proteins were transferred electrophoretically onto polyvinylidene difluoride membranes and were blocked with 5% nonfat dry milk and subsequently incubated overnight at 4° C with primary polyclonal iNOS, polyclonal nNOS, or polyclonal ecNOS antibodies. Lysates of cytokine-activated macrophages, pituitary cells, and endothelial cells were used as positive controls for iNOS, nNOS, and ecNOS proteins, respectively (provided by Transduction Laboratories Inc.). Specific proteins were detected using horseradish peroxidase-conjugated anti-rabbit secondary antibodies and ECL reagents (Amersham Corp.). The blots were scanned with an imaging densitometer (Model GS700, Bio-Rad Inc., 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 Inc.) were used as markers. The membranes were stained with the silver stain and scanned to verify that equal amounts of proteins were loaded among different lanes.

Muscle NOS Activity

Aliquots from muscle homogenates (100-200 µg) were incubated within 50 mmol/L HEPES (pH 7.4) with 100 mmol/L [³H]arginine (50 Ci/mmol), 120 µmol/L NADPH, 60 mmol/L L-valine, 12 mmol/L L-citrulline, 1.2 mmol/L MgCl₂, 0.2 mmol/L CaCl₂, 10 µg/ml calmodulin, 3 µmol/L BH4, 1 µmol/L FAD, and 1 µmol/L FMN. The reaction was carried out for 1 h at 37° C with and without 2 mmol/L N^G-nitro-L-arginine methyl ester (L-NAME, NOS inhibitor) and terminated by adding 2 ml of 20 mmol/L HEPES (pH 5.5) containing 2 mmol/L EDTA. To measure Ca²⁺-independent NOS activity, the reaction was repeated in the presence of 1 mmol/L EDGTA. Samples were then applied to 1-ml columns of Dowex AG50W-X8 (Na⁺ form), which were eluted with 2 ml of water. [³H]Citrulline was quantified by liquid scintillation spectroscopy of 4.0 ml flow through. Specific NOS activity was obtained by subtracting [³H]citrulline formed in the presence of L-NAME from the total [³H]citrulline recovered. Calcium-independent NOS activity was determined by evaluating [³H]citrulline formation in the absence of calcium (the presence of EGTA).

Diaphragmatic Strip Preparation

This preparation was used to prepare the diaphragm for either contractility measurement or assessment of sarcolemmal integrity. Diaphragms from the four groups of mice 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 that had 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 (2 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).

Force-frequency Measurements

Muscle strips were stimulated electrically at constant currents via platinum electrodes mounted in the muscle chamber and connected to a square wave pulse stimulator (Grass Instruments, Model S48). After an equilibration period of 30 min (temperature of 22-25° C), the organ bath temperature was increased to 35° C and the maximum current necessary to elicit maximum 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 maximum isometric muscle force was generated in response to supramaximal stimulation (current of 300-350 mA, 120-Hz frequency). Muscle contractility was evaluated by stimulating the muscle at 10, 20, 30, 50, 100, and 120 Hz while maintaining constant supramaximal current and stimulation duration (600 ms). 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 (centimeters) and weight (grams) 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. Peak muscle force in N/cm² was measured for each contraction within the force-frequency curve.

Assessment of Sarcolemmal Injury

Isolated diaphragmatic strips obtained from the four animal groups were prepared as mentioned above and were assigned for the measurements of sarcolemmal injury under resting (unstimulated protocol) conditions or after a short period of intermittent titanic stimulations (stimulated protocol). In both groups, the fluorescent molecular tracer, Procion orange 14 (Sigma Inc.), was added to the Krebs (0.15% wt/vol solution) for identification of fibers undergoing sarcolemmal injury. The total continuous exposure of the strips to the bath solution containing Procion orange was at least 90 min for all of the groups. In the unstimulated protocol, muscle strips were rinsed off Procion orange for 5 min in normal Krebs solution and then quick-frozen in isopentane precooled with liquid nitrogen, and preserved at 80° C. In the stimulated protocol, muscle strips were stimulated at 50 Hz, 300 ms duration, for a total period of 3 min. When the stimulation protocols were completed, the muscle strips were rinsed as in the unstimulated protocol and frozen. Serial sections (10 µm thick) were cut in the transverse plane of both stimulated and unstimulated muscle samples with a cryostat microtome (Leica Cryocut 1800, Heidelberg, Germany). To assess the amount of sarcolemmal injury, transverse frozen sections of rat diaphragm were randomly selected and photographed using epifluorescence microscopy (Nikon E600, Japan). Using the FITC filter setting and a magnification level of ×100, 10-15 microscopic fields (minimum of 1000 fibers) per muscle were analyzed by using the MetaMorph (Universal Imaging Corporation, PA) imaging software package. Muscle fibers demonstrating a clear increase in cytoplasmic fluorescence (i.e., fibers containing the Procion Orange 14 tracer dye) were counted, and the percentage of dye-positive fibers on each diaphragm section was determined. Areas with sectioning artifacts (folds, tears, etc.) were avoided, and diaphragmatic edges of the sections were also excluded to avoid areas damaged by the muscle dissection. The counting was done following a double-blind protocol.

Data Analysis

The data on NOS protein expression, NOS activity, muscle force, and sarcolemmal injury are presented as means ± SEM. Differences between various groups were compared by two-way analysis of variance for repeated measures. Any differences detected were evaluated post hoc by the Student Neuman-Keuls test. p < 0.05 was considered significant.

	RESULTS

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Figure 1 illustrates representative immunoblots of diaphragmatic protein samples probed with antibodies selective to nNOS, iNOS, and ecNOS isoforms. Prominent nNOS protein bands were detected in WT-C and WT-LPS groups, whereas no detectable nNOS protein bands were observed in nNOS^/ mice (Figure 1). By comparison, very weak iNOS protein levels were detected in the control groups of both WT and nNOS^/ mice. Injection of LPS elicited a significant induction of iNOS protein both in WT and nNOS^/ mice, however, the intensity of iNOS expression, as evaluated by optical density of iNOS protein (n = 6 in each group), was significantly lower in the nNOS^/ mice compared with WT mice (Figures 1 and 2). No differences were detected in the expression of ecNOS protein among the four groups of animals (Figures 1 and 2).

View larger version (32K): [in this window] [in a new window]   Figure 1.   Representative immunoblot of diaphragmatic protein samples probed with anti-nNOS (top), iNOS (middle), and ecNOS (bottom) antibodies. Notice the absence of nNOS protein in the nNOS/ mice samples. Also note the weak expression of iNOS protein in WT-C and nNOS/-C groups, whereas more prominent iNOS protein expression was evident in WT-LPS and nNOS/-LPS groups.

View larger version (32K): [in this window] [in a new window]   Figure 2.   Mean values of iNOS and ecNOS protein densities after LPS injection in WT (n = 6) and nNOS/ (n = 6) mice. +p < 0.05 compared with WT mice.

Figure 3 shows the changes in diaphragmatic NOS activity in the four groups of animals. Injection of LPS in WT mice resulted in augmentation of total muscle NOS activity from 2.09 to 3.17 pmol/mg/min. This rise was due to a substantial rise in Ca²⁺-independent NOS activity, which was not detectable in the control WT animals but reached 1.91 pmol/mg/min in WT-LPS animals (p < 0.01 compared with control animals, Figure 3). Total muscle NOS activity in both nNOS^/-C and nNOS^/-LPS animals was much lower than that of WT animals and showed no detectable levels of Ca²⁺-independent activity (p < 0.01 compared with WT animals, Figure 3).

View larger version (16K): [in this window] [in a new window]   Figure 3.   Diaphragmatic NOS activity in the four groups of animals. Notice the significant rise in Ca2+-independent NOS activity in response to LPS injection in WT animals. Also note that both nNOS/-C and nNOS/-LPS mice developed much lower levels of NOS activity compared with WT mice.

Figure 4 illustrates the changes in diaphragmatic contractility as measured by the force-frequency relationship in the four groups of animals (n = 6 in each group). Injection of LPS in the WT animals produced a significant decline in diaphragmatic force at frequencies higher than 20 Hz (p < 0.05 compared with the control group). Similarly, LPS injection was associated with a significant decline in diaphragmatic force in nNOS^/ mice (p < 0.05 compared with the control nNOS^/ group). However, LPS-induced reduction in diaphragmatic force at frequencies higher than 30 Hz was more severe in nNOS^/ mice compared with WT animals (p < 0.05, Figure 4).

View larger version (19K): [in this window] [in a new window]   Figure 4.   Changes in diaphragmatic contractility in the four groups of animals. *p < 0.05 compared with control groups. +p < 0.05 compared with corresponding WT groups. Note the significant decline in diaphragmatic force in the nNOS/-LPS group compared with the WT-LPS group.

Representative micrographs illustrating histological sections of diaphragmatic samples isolated from the four groups of animals, artificially stimulated in vitro and then stained with Procion orange dye, are shown in Figure 5. Sarcolemmal damage is indicated by positive dye staining inside the sarcoplasm. Figure 6 shows the mean values of the percentage of injured fibers in the four groups of animals under unstimulated and stimulated conditions. Under unstimulated conditions, the percentage of injured fibers was less than 5% of total muscle fibers in the four groups of animals and no significant differences were noticed among the four groups. By comparison, stimulation of diaphragmatic strips of saline-injected WT animals was associated with a substantial degree of muscle injury with positive fibers constituting about 27% of total fibers. After 12 h of LPS injection in WT mice, stimulation of diaphragmatic strips elicited a significantly greater degree of muscle fiber injury than that observed after saline injection (p < 0.05). Similarly, the extent of muscle fiber injury in nNOS^/ mice was significantly higher after LPS injection than after saline injection (p < 0.05, Figure 6). However, there were no significant differences between nNOS^/ and WT mice in the degree of muscle fiber injury after either saline or LPS injection (Figure 6).

View larger version (59K): [in this window] [in a new window]   Figure 5.   Transverse sections of diaphragm treated with the fluorescent dye Procion Orange 14. The fluorescent fibers indicate sarcolemma lesions in WT-C (A), nNOS/-C (B), WT-LPS (C), and nNOS/-LPS (D) groups. Note the significant rise in the percentage of injured fibers after LPS injection in both WT and nNOS/ mice. Bars indicate 100 µM.

View larger version (26K): [in this window] [in a new window]   Figure 6.   The extent of muscle fiber injury (as percentage of total fibers) in the four groups of animals measured under resting (unstimulated) conditions and after 3 min of intermittent artificial stimulation (stimulated). *p < 0.05 compared with corresponding control groups. Note the significant rise in the degree of sarcolemmal injury following muscle stimulation. Also note that the absence of nNOS had no significant effect on LPS-induced sarcolemmal injury.

	DISCUSSION

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The main findings of this study are as follows (1) LPS injection elicited a significant decline in diaphragmatic contractility that was more severe in nNOS^/ mice compared with WT mice. (2) LPS injection was also associated with a significant increase in stimulation-induced diaphragmatic sarcolemmal injury compared with normal animals, however, the extent of LPS-induced sarcolemmal injury was similar among nNOS^/ and WT mice. (3) The induction of iNOS expression in the diaphragm in response to LPS injection was weaker in nNOS^/ mice compared with WT mice. (4) Muscle NOS activity in both control and LPS-injected nNOS^/ mice was significantly lower than that measured in WT animals.

Role of NO in Muscle Contractility

Under normal conditions, NO is synthesized inside skeletal muscle fibers by the nNOS isoform, which is localized at the sarcolemma and associates with the dystrophin complex (10). The exact functional importance of endogenous NO synthesis in regulating normal muscle function is still being investigated. Inhibition of endogenous NO synthesis in mature or embryonic muscles results in a significant elevation of submaximal muscle force indicating that basal muscle NOS activity exerts an inhibitory influence on contractile performance (11). The mechanisms through which NO modulates muscle contractility are not fully understood but several sites are likely to be influenced by NO including neuromuscular transmission, sarcoplasmic reticulum Ca²⁺ kinetics, and the activities of creatine kinase and myosin ATPase.

Our current results indicate that the absence of nNOS in nNOS^/ mice resulted in worsening of the LPS-induced decline in diaphragmatic contractility. These results and our recently published study in which LPS-induced reduction in diaphragmatic contractility was more severe in iNOS^/ mice compared with WT mice (8) suggest that both the iNOS and nNOS isoforms play protective roles in attenuating LPS- induced muscle dysfunction. An interesting finding in our study is that the absence of nNOS worsened muscle contractility at frequencies of stimulation higher than 30 Hz (Figure 4). This observation can be attributed to the fact that nNOS is expressed mainly in fast-twitch (type II) muscle fibers (11). Thus, absence of nNOS will likely be associated with a preferential reduction in high-frequency force output. There are several possible pathways through which the nNOS isoform may protect muscle function in septic animals. First, NO enhances Ca²⁺ release from the sarcoplasmic reticulum by activating ryanodine receptors (12). NO can also protect these receptors from the oxidative effects of reactive oxygen species and, hence, causes a rise in intracellular Ca²⁺ levels (13). Recent evidence also indicates that NO enhances tetanic cytosolic Ca²⁺ concentrations through voltage-sensitive Ca²⁺ channels (14). Second, NO enhances fuel oxidation in skeletal muscle fibers through an increase in glucose utilization particularly in response to muscle contraction (15). The enhancement of fuel utilization by NO is particularly important in counteracting the influence of sepsis on skeletal muscle glucose uptake. Third, it is possible that the protective effect of nNOS in LPS-injected animals may be mediated in part through the promotion of iNOS expression. In the current study, both the expression of iNOS protein and Ca²⁺-independent NOS activity were much lower in nNOS^/-LPS mice compared with WT-LPS mice. We have recently demonstrated that mice deficient in iNOS develop a much more severe form of LPS-induced muscle contractile dysfunction than wild-type animals (8). Accordingly, lower levels of iNOS expression in nNOS^/-LPS compared with WT mice are expected to worsen muscle contractility. Fourth, it has been established that ventilatory muscle contractile dysfunction in sepsis is mediated in part by an increased rate of ROS (16, 17). This notion is confirmed by the observations that infusion of ROS scavengers and antioxidants such as N-acetylcysteine or superoxide dismutase results in improvement of ventilatory muscle contractility in septic animals (16). It has been argued that endogenous NO synthesis inside skeletal muscle fiber may reduce ROS levels by directly scavenging superoxide anions and, therefore, reduces the conversion of this radical to more reactive radicals such as H₂O₂ and HO. NO is also capable of reducing ROS-generation by inhibiting the activities of ROS-generating enzymes such as NADPH oxidase and xanthine oxidase (18, 19). Finally, NO is known to induce the expression of heme oxygenase 1 in skeletal muscle fiber (20). Heme oxygenase 1 exerts potent antioxidant effects, which are mediated by its products, carbon monoxide and bilirubin. We should emphasize, however, that in vivo NO chemistry is complex and many studies have pointed out that NO can function as both an antioxidant molecule or prooxidant agent depending on various local factors such as the concentration of NO, the availability of NOS substrate and cofactors, and the local levels of ROS (for review, see Wink and Mitchell [21]).

Interactions between nNOS and iNOS Expressions

Little is known about the nature and the mechanisms of interactions between the expression of various NOS isoforms particularly during the course of sepsis or septic shock. We reported in our previous study that the absence of iNOS protein in iNOS^/ mice was associated with a significant induction in muscle nNOS expression in septic animals indicating that iNOS activity exerts an inhibitory influence on nNOS expression (8). We found in the current study, however, that nNOS-deficient mice develop a significantly lesser degree of iNOS protein expression and iNOS activity in the diaphragm in response to LPS injection than WT mice (Figures 1, 2, and 3). The exact mechanisms responsible for this observation are not clear. We propose NO and NO-related species activate guanine nucleotide-binding protein p21^ras and mitogen-activated protein (MAP) kinases, which, in turn, lead to downstream signaling events such as nuclear translocation of NF-B (22). This pathway, which favors a positive influence of NO on iNOS induction, can explain the lower degree of iNOS induction in LPS-injected nNOS^/ mice as compared with WT mice. We should point out that in the current study we did not assess the colocalization of iNOS and nNOS proteins in LPS-injected mice. Thus, we are uncertain whether reduction in total muscle iNOS expression in nNOS^/-LPS mice was due to selective attenuation of iNOS induction inside type II fibers (main sites of nNOS expression in normal skeletal muscles) or due to lower levels of iNOS expression in other sites inside skeletal muscles such as infiltrating inflammatory cells, which are known to express iNOS in septic rats (3). Clearly, the nature of the interactions between nNOS and iNOS isoforms and the mechanisms behind this interaction need to be thoroughly investigated.

Sarcolemmal Injury

Skeletal muscle sarcolemmal injury has been studied extensively in limb muscles and has been attributed mainly to mechanical disruption of the cell membrane associated with eccentric contractions. The development of sarcolemmal injury in the ventilatory muscles has recently been documented in conscious dogs by Zhu and coworkers (23) who reported an increase in the percentage of injured fibers from 0.3% to 7.6% of total muscle fibers following mild resistive loading. In rat muscles, Zhu and coworkers (24) recently indicated that the degree of sarcolemmal injury correlates significantly with total tension time index generated by artificial in vitro stimulation. Our observation that the levels of sarcolemmal injury in stimulated muscles were substantially greater than those of unstimulated muscle is in accordance with that of Zhu and coworkers (23, 24).

We have recently demonstrated that sepsis induced by either LPS injection or cecal ligation in rats elicited sarcolemmal damage that was more severe in the ventilatory than in limb muscles and coincided with iNOS induction in these muscles (7). The observation that the degree of sarcolemmal injury in septic animals was significantly reduced when NOS inhibitors were infused systemically led to the conclusion that NO, synthesized mainly by the iNOS isoform, plays a role in maintaining of sarcolemmal integrity (7). The current study suggests, however, that the absence of nNOS protein did not alter the degree of sepsis-induced sarcolemmal injury in the diaphragm (Figures 5 and 6). The reasons behind these contradictory conclusions regarding the role of NO in the pathogenesis of sarcolemmal injury in septic animals are not clear. We speculate, however, that dissimilarities in the rate of skeletal muscle NO synthesis in normal and septic rats versus mice and methodological differences between the two studies are involved. For instance, Lin and coworkers (7) evaluated the role of NO by infusing NOS inhibitors systemically, a procedure that is likely to be associated with secondary effects on muscle sarcolemmal integrity as a result of changes in muscle blood flow and O₂ consumption.

Our observation of a similar degree of LPS-induced sarcolemmal injury between nNOS^/ and WT mice is rather surprising as nNOS possesses many features that support the possibility of a protective role against the development of sarcolemma injury. These features include its localization in close proximity to the sarcolemma and its direct association with the dystrophin complex (10), a major player in the maintenance of sarcolemmal integrity. In addition, the observation that type I fibers (devoid of or having very low levels of nNOS protein) are more susceptible than type II fibers (rich in nNOS) to resistive loading-induced sarcolemmal injury suggests that nNOS may play a protective role in maintaining sarcolemmal injury (23). However, our study strongly supports the conclusion that nNOS does not play a major role in the pathogenesis of "mechanical stress-induced sarcolemmal injury." It should be emphasized that the exact mechanisms responsible for sarcolemmal injury in septic animals remain under investigation. Important factors other than NO, which are likely to be involved, include changes in intracellular Ca²⁺ levels, enhanced lipid peroxidation, and membrane protein oxidations as a result of reactive oxygen species production inside muscle fibers or oxidants derived from infiltrating leukocytes.

In summary, our results indicate that LPS injection elicits different effects on diaphragmatic NOS protein expression and contractility in WT and nNOS^/ mice. Maximal force and iNOS induction were lower after LPS injection in nNOS^/ mice compared with WT mice, however, differences in nNOS genotype did not alter the degree of LPS-induced sarcolemmal injury in the diaphragm. These results suggest that the nNOS isoform may play a protective role in attenuating the negative effects of LPS on diaphragmatic contractility but not on sarcolemmal integrity.