Journal of Applied Physiology
Vol. 81, No. 6, pp. 2415-2420, December 1996
EXERCISE AND MUSCLE

Expression of the inducible nitric oxide synthase gene in diaphragm and skeletal muscle

Marita Thompson, Lisa Becker, Debbie Bryant, Gary Williams, Daniel Levin, Linda Margraf, and Brett P. Giroir

Laboratories for Molecular Biology and Physiology Research, Department of Pediatrics, and Department of Pathology, The University of Texas Southwestern Medical Center, Dallas, Texas 75235-9063

ABSTRACT

INTRODUCTION

METHODS

RESULTS

DISCUSSION

FOOTNOTES

REFERENCES

ABSTRACT

Thompson, Marita, Lisa Becker, Debbie Bryant, Gary Williams, Daniel Levin, Linda Margraf, and Brett P. Giroir. Expression of the inducible nitric oxide synthase gene in diaphragm and skeletal muscle. J. Appl. Physiol. 81(6): 2415-2420, 1996.Nitric oxide (NO) is a pluripotent molecule that can be secreted by skeletal muscle through the activity of the neuronal constitutive isoform of NO synthase. To determine whether skeletal muscle and diaphragm might also express the macrophage-inducible form of NO synthase (iNOS) during provocative states, we examined tissue from mice at serial times after intravenous administration of Escherichia coli endotoxin. In these studies, iNOS mRNA was strongly expressed in the diaphragm and skeletal muscle of mice 4 h after intravenous endotoxin and was significantly diminished by 8 h after challenge. Induction of iNOS mRNA was followed by expression of iNOS immunoreactive protein on Western immunoblots. Increased iNOS activity was demonstrated by conversion of arginine to citrulline. Immunochemical analysis of diaphragmatic explants exposed to endotoxin in vitro revealed specific iNOS staining in myocytes, in addition to macrophages and endothelium. These results may be important in understanding the pathogenesis of respiratory pump failure during septic shock, as well as skeletal muscle injury during inflammation or metabolic stress.

endotoxin; cytokines; septic shock

INTRODUCTION

NITRIC OXIDE (NO) is a multifunctional molecule that participates in regulation of vasomotor tone, microbiological defense, neural transmission, and intracellular signal transduction (28). NO is produced during the oxidation of arginine to citrulline by constitutive and inducible isoforms of the enzyme NO synthase (NOS) (30, 31). Recently, Balon and Nadler (3) demonstrated NO release from incubated normal rat extensor digitorum longus muscle preparations but did not definitively determine whether NO was secreted by skeletal myocytes or other cells in their preparations, e.g., endothelial cells (37). Simultaneously with these observations, Kobzik and co-workers (24) documented the presence of the neuronal constitutive isoform of NOS (ncNOS) in rat skeletal muscle by immunocytochemistry. These results substantiated earlier work by Nakane et al. (29), who demonstrated the presence of ncNOS mRNA in human skeletal muscle. These data have fueled speculations that NO may play an important role in the physiology of skeletal muscle contraction or metabolism (38).

Inducible NOS (iNOS) has also been demonstrated in a number of cell types, including macrophages, hepatocytes, vascular smooth muscle cells, epithelial cells, and endothelial cells (18, 26, 27, 35). Recently, we have described the induction of iNOS in skeletal muscle myotubes and myoblasts of the C2C12 lineage in vitro (41). In C2C12 cells, iNOS was induced by combinations of cytokines that included interferon- and yielded biosynthesis of NO in quantities equivalent to these produced by macrophages under similar conditions.

It currently remains unknown whether the iNOS gene is also expressed by skeletal muscle in vivo. Given recent data suggesting the importance of NO produced by ncNOS in normal skeletal muscle, we determined whether iNOS might be induced in skeletal muscle and diaphragm during inflammatory conditions in vivo. Because NO has been shown to depress skeletal muscle contractility in vitro (24), expression of iNOS by skeletal muscle during endotoxemia may elucidate the pathogensis of respiratory pump failure and muscle catabolism during septic shock.

METHODS

Animals. Six- to eight-week-old male C3H/HeN mice (Sasco) were housed in a thermal- and light-controlled animal facility and allowed free access to chow and water. Experimental and control animals were injected intravenously through the tail vein with either 50 µg of Escherichia coli O111:B4 lipopolysaccharide (LPS; Sigma Chemical) or an equivalent volume of sterile normal saline. Animals were monitored by using a rodent distress assessment that monitored body weight, appearance, clinical signs, and unprovoked behavior as measures of distress. Animals were killed at the indicated times after CO₂ narcosis followed by cervical dislocation.

RNA purification and Northern blots. Tissues were carefully excised from the killed animals and rinsed thoroughly in normal saline. Samples were obtained from the cardiac ventricles (atria excluded), each hemidiaphragm, and quadriceps skeletal muscles. Total RNA was purified by a modification of the method of Chomczynski and Sacchi (11) by using RNAzol B (Biotecz Labs, Houston, TX). The RNA was quantitated by absorbance at 260 nm, and 10 µg of total RNA were resolved on a 1.2% agarose gel as previously described. The iNOS probe consisted of a 765-bp fragment complementary to the 5 end of the murine macrophage iNOS cDNA (41). Experiments were independently repeated seven times, with groups as indicated in RESULTS.

Western immunoblots. Tissues were immediately harvested from the killed animals. An equivalent amount of tissue (depending on the tissue examined, between 50 and 150 mg) was homogenized in 800 µl of ice-cold grind buffer containing 50 mM tris(hydroxymethyl)aminomethane (Tris), pH 7.4, 1 mM EDTA, 0.1 mM dithiothreitol (DTT), 4 µM bestatin, 1 mM N-p-tosyl-L-lysine chromomethyl ketone (TLCK), 1 mM phenylmethylsulfonyl fluoride (PMSF), 20 mM 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS), and 10 µg/ml each of leupeptin, pepstatin A, and aprotonin. The sample was then centrifuged for 10 min at 10,000 g at 4°C, and supernatant was collected. To this supernatant were then added 50 µl of preswollen 2-5 ADP-sepharose-4B (Pharmacia), which was then gently rocked for 1 h at 4°C. The preswollen 2-5ADP-sepharose-4B was composed of one-third volume 2-5-sepharose-4B and two-thirds volume grind buffer. After an additional centrifugation and after discarding of the supernate, 100 µl of an elution buffer consisting of 1× homogenization buffer, 10 mM NADP, and 10% vol/vol glycerol were added to the conjugated ADP-sepharose. After incubation for 15 min on ice, supernatant was removed after centrifugation. Forty microliters of this supernatant were added to an equal volume of gel-loading buffer (50 mM Tris, pH 6.8, 100 mM DTT, 2% sodium dodecyl sulfate, 0.1% bromphenol blue, and 10% glycerol), boiled for 10 min at 100°C, resolved on a 6% polyacrylamide gel, then transferred to a nitrocellulose membrane. The membrane was blocked overnight at 4°C in 5% dried milk dissolved in Tris-buffered saline-Tween 20 (TBS-T) (20 mM Tris, pH 7.6, 137 mM NaCl, 0.1% Tween 20). The membrane was then washed in TBS-T and incubated for 1 h with a monoclonal anti-mouse iNOS antibody (Transduction Laboratories) used at 1:500 dilution. After two additional washes in TBS-T, the membrane was incubated for 1 h at room temperature with a 1:8,000 dilution of sheep anti-mouse immunoglobulin G horseradish peroxidase-linked antibody (Amersham). After an additional wash in TBS-T, the membrane was exposed to the mixture of luminol plus hydrogen peroxide under alkaline conditions, and the strength of the resulting chemiluminescent reaction was registered by autoradiography. Experiments were done independently three times, with groups as indicated in RESULTS.

iNOS activity. iNOS activity was determined by measuring the conversion of [³H]arginine to [³H]citrulline. Tissue was carefully excised from the killed animals and homogenized in 1,000 µl of ice-cold buffer containing 50 mM Tris, pH7.4, 1 mM EDTA, 0.1 mM DTT, 4 µM bestatin, 1 mM TLCK, 1 mM PMSF, 20 mM CHAPS, and 10 µg/ml each of leupeptin, pepstatin A, and aprotonin. The sample was then centrifuged for 5 min at 10,000 g at 4°C, and the supernatant was collected. Fifty microliters of the supernatant were then added to 50 µl of buffer containing 35 mM N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid (HEPES), 4 mM B-NADPH, 20 µM tetrahydrobiopterin, 20 µM flavin adenine dinucleotide, 20 µM flavin mononucleotide, 1.0 mM CaCl₂, 30 nM calmodulin, 4 µM cold L-arginine, 4 mM ethylene glycol-bis(-aminoethyl ether)-N,N,N,N-tetraacetic acid (EGTA), and 2.0 µCi/ml L-[³H]arginine. It was incubated at 37°C for 60 min, then the assay was terminated by the addition of buffer containing 40 mM HEPES, pH 5.5, 2 mM EDTA, and 2 mM EGTA. The samples were then applied to 1-ml Dowex AG50WX-8 (Tris form) columns and eluted with 1 ml of the 40 mM HEPES buffer. The effluent was collected into scintillation vials and quantified by liquid scintillation spectroscopy. Protein content was determined by using the Bradford technique (Bio-Rad) with albumin as a standard. Results are expressed as units per microgram protein = picomoles per minute per microgram protein. Experiments were performed three independent times, with groups as indicated in RESULTS.

Immunochemistry. Explants from normal mouse diaphragm (~3 × 3 mm) were incubated for 24 h in Dulbecco's modified Eagle medium (Sigma Chemical) supplemented with 5% heat-inactivated fetal bovine serum (GIBCO BRL) and 4% penicillin-streptomycin solution (GIBCO BRL). Cultures were maintained in a humidified atmosphere of 5% CO₂ in an incubator at 37°C. Explants were either exposed or not exposed to 1 µg/ml LPS (E. coli O111:B4, Sigma Chemical) for 24 h. After incubation, tissue was fixed in 100% ethanol and routinely processed for light microscopy and embedded in paraffin. Four-micrometer-thick sections were deparaffinized and incubated overnight at room temperature with a 1:50 dilution of mouse monoclonal anti-iNOS antiserum (Transduction Labs, Lexington, KY). Immunoreactivity was detected by using streptavidin-biotin methodology with commercially available kits, according to the manufacturer's specifications (Cell Marque, Austin, TX). Negative controls, consisting of sections incubated in buffer alone, were run concurrently.

Statistical analysis. Values for mRNA densitometry and [³H]arginine are expressed as means ± SE. The statistical significance was determined by one-way analysis of variance with Bonferroni correction. Statistical significance was set at P < 0.05.

RESULTS

Animals that had received intravenous endotoxin showed moderate distress scores based on anorexia, piloerection, lethargy, and 3% weight loss. These animals demonstrated the induction of iNOS mRNA in diaphragm and skeletal muscle, which peaked at 4 h and was significantly diminished by 8 h after challenge. Levels of iNOS mRNA in the diaphragm were comparable to those induced in the myocardium, whereas the levels induced in skeletal muscle were higher compared with controls (Fig. 1). Densitometric analysis (Fig. 1) demonstrated a fivefold increase in the diaphragm and a sevenfold increase in the quadriceps skeletal muscle, compared with control levels. iNOS mRNA was not demonstrated in sheep pulmonary artery endothelium. Because iNOS gene expression is significantly regulated at the level of translation, we next verified that an induction of iNOS mRNA was mirrored by an increase in the tissue level of iNOS immunoreactive protein. Western immunoblots of ADP-sepharose-fractionated protein extracts (Fig. 2) demonstrated nondetectable or only minimally detectable iNOS immunoreactive bands under control conditions; however, after endotoxin challenge, the iNOS bands, as detected by monoclonal antibody to mouse macrophage iNOS, were significantly increased at both 8 and 12 h after challenge, in both the diaphragm and skeletal muscle as well as in the heart. We found that iNOS protein was present, and, in addition, that iNOS activity (Fig. 2) was also present and increased after LPS administration. The conversion of arginine to citrulline not only documented the presence of iNOS activity but also quantified its activity on a microgram of protein basis. This quantification is important, as it is difficult to quantitate the protein used in the ADP-sepharose extract-based Western blots. The ADP-sepharose binds selectively to iNOS to enrich the iNOS content, which results in specimens that are semiqualitative. Activity expressed as units per microgram of protein was significantly increased from control levels at 8 and 12 h after LPS administration. iNOS activity levels were over fivefold higher in both the diaphragm and skeletal muscle at 8 h and remained elevated above control levels at 12 h post LPS administration. iNOS activity was fully inhibited by the addition of N-nitro-L-arginine methyl ester in all tissue types (results not shown).

1

1

Although it is unlikely that endothelial cells or infiltrating macrophages could account for the observed significant increase in iNOS mRNA and immunoreactive protein, we next sought to localize iNOS immunoreactive protein to the myocytes themselves. Sections of diaphragm from LPS-treated mice stained immunochemically for iNOS showed moderate immunoreactivity within the endothelium of blood vessels within the muscle. Rare isolated intermuscular cells, morphologically consistent with macrophages, were strongly immunoreactive (arrow). Muscle fibers were negative (Fig. 3A). The diaphragm from control animals showed less endothelial reactivity and no evidence of macrophage or myocyte iNOS production (Fig. 3B). Because the lack of myocyte immunostaining could be explained by the insensitivity of immunochemistry compared with Northern and Western analyses, we attempted to increase the levels of iNOS expression by incubating diaphragm explants in vitro with or without the presence of bacterial LPS. Microscopic sections of the diaphragm explants showed cytoplasmic immunoreactivity for iNOS within myocytes. Low levels of iNOS expression were present in control explants incubated for 24 h (Fig. 3D) compared with nonincubated explants (Fig. 3E). iNOS immunoreactivity (brown stain) was significantly increased in myocytes from explants exposed to LPS for 24 h (Fig. 3C).

DISCUSSION

We have previously reported the induction and expression of the macrophage-type iNOS gene in C2C12 skeletal myoblasts and myotubes in vitro (41). To our knowledge, the experiments reported here are the first to demonstrate induction of iNOS mRNA, iNOS protein, and iNOS activity in skeletal muscle and diaphragm in vivo. Induction of immunoreactive iNOS protein was demonstrated in diaphragm in vitro, although not in vivo. This is consistent with previous iNOS immunochemical findings. Despite the fact that several investigators have convincingly demonstrated NO production from cardiac myocytes (9), cardiac myocytes have not been immunoreactive for iNOS. Buttery et al. (9) immunostained for iNOS protein in LPS-exposed rats and were unable to demonstrate iNOS staining in cardiac myocytes, although staining of macrophages was noted. Yet, Balligand et al. (2) demonstrated NO release from single cytokine-stimulated rat cardiac myocytes. The lack of iNOS immunostaining in the cardiac myocytes was thought to be secondary to lack of sensitivity of the immunochemical technique to lower levels of iNOS protein.

The precise role of iNOS expression in striated muscle during endotoxemia or other provocative states is yet unknown; nonetheless, it is possible that iNOS expression may be detrimental and lead to respiratory muscle dysfunction; alternately, iNOS expression may be adaptive in that production of NO may lead to enhanced oxygen and substrate delivery to vital tissues during prolonged periods of disease and/or physiological stress. Thus, since it not known whether increased NO production plays a pathophysiological, compensatory, or incidental role in septic shock, the relevance of these findings to human septic shock is not yet determined.

During septic shock of gram-negative and gram-positive etiologies, diaphragmatic function is abnormal. In a canine endotoxic shock model, Hussain et al. (21) demonstrated that death resulted not from cardiac failure but from respiratory compromise and, ultimately, respiratory failure. These and other investigators have since determined that sepsis causes diaphragmatic dysfunction evidenced by diminished diaphragmatic contractility and reduced endurance (5, 20). Dysfunction of the diaphragm can be induced in vitro by LPS-activated monocyte supernatant but is not the result of a direct and immediate effect of tumor necrosis factor- or endotoxin (12, 40). It is possible that NO produced by the diaphragm, induced by endotoxin and cytokines, may account for the diaphragmatic dysfunction witnessed during sepsis. The recent report by Kobzik et al. (24) documented that production of NO by diaphragm via ncNOS diminished contractility, whereas inhibitors of NOS activity and guanosine 3,5-cyclic monophosphate augmented contractile function. Precedent for a contractility depressant effect of NO is also found in the literature investigating sepsis-induced myocardial dysfunction. Although these data remain controversial (39), several investigators have demonstrated that NO causes contractile dysfunction in both cardiac myocytes and papillary muscles (1, 7, 8, 15, 16, 23). In addition, intracoronary infusion of the NO donor nitroprusside into nonseptic humans resulted in reduced left ventricular pressure development and a left ventricular relaxation hastening effect (32).

Detrimental effects of NO on muscle function could be mediated directly through the stimulation of soluble guanylate cyclase, indirectly through guanosine 3,5-cyclic monophosphate-dependent protein kinases, or by interactions of NO with transcription factors (25, 28). NO has also been demonstrated to reversibly inhibit cytochrome c in skeletal muscle mitochondrial respiration. This inhibition could decrease ATP production, ultimately resulting in respiratory muscle dysfunction (12). Additionally, it is known that skeletal muscles produce reactive oxygen radicals during stress (33, 34); reactive oxygen radicals could combine with NO to form peroxynitrite and, thereby, lead to cell injury or death (4, 17). Consistent with this speculation is a recent report that documented skeletal muscle edema and fiber necrosis during a porcine model of septic shock (19).

Alternately, the induction of NOS in diaphragm and skeletal muscle may be adaptive during certain conditions in that prolonged exertional or metabolic stress would increase NO production and thereby improve organ blood flow. This may be particularly important during endotoxic shock, during which time diaphragmatic oxygen extraction is impaired, and a pathological oxygen supply-demand relationship may exist (22). In regard to nondiaphragm skeletal muscle, a number of cytokines, including interleukin-1, tumor necrosis factor-, and interferon-, are known to be elevated in skeletal muscle and/or in serum after prolonged vigorous exercise (6, 10, 14). NO of skeletal muscle origin may also function to enhance local oxygen and substrate delivery and, thereby, be adaptive during such metabolic or exertional stresses.

In 1991, Salter and colleagues (36) surveyed biochemical evidence of NOS activity in multiple tissues from three species. These investigators reported the presence of calcium-independent NOS activity in the rat diaphragm, but not in rat, rabbit, or guinea pig skeletal muscle, 6 h after intraperitoneal endotoxin challenge. The present data confirm the inferences of Salter and co-workers that calcium-independent iNOS is induced in the diaphragm and extend their observations to include nonrespiratory skeletal muscle. The addition of EGTA as a calcium chelator to our assay ensured measurement of calcium-independent iNOS activity, since brain cell NOS and ecNOS are calcium-dependent enzymes. There are several possible explanations for the detection of iNOS in our system and not in that of Salter et al. First, our experiments were done on mice, and therefore a species-specific difference may account for the discrepancy. Which species most closely approximates human iNOS responses is yet unknown. In addition, the current methodology was designed to directly detect the iNOS gene products (both mRNA and protein), in contrast to detection of citrulline as indirect evidence of NOS induction and/or activity. Alternatively, although we were able to demonstrate the induction of iNOS after both intraperitoneal and intravenous endotoxin administration, induction of iNOS in nondiaphragmatic skeletal muscle was much greater with intravenous endotoxin administration, even when the intravenous doses of LPS were 20% of those administered intraperitoneally (data not shown). We demonstrated an increase in iNOS protein production and activity. It is also possible that NOS cofactors levels, such as tetrahydrobiopterin, may be affected by LPS, which may also influence iNOS activity.

The present study supports the concept that NO may be produced by numerous cell types within tissues of diverse origins. The net beneficial or detrimental effects of this pluripotent molecule may depend on the particular circumstances of its production and the physiological state of the organism in which it is produced.

FOOTNOTES

Address for reprint requests: B. P. Giroir, Univ. of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75235-9063 (E-mail: Giroir{at}UTSW.SWMED.EDU).

Received 16 March 1995; accepted in final form 26 July 1996.

REFERENCES

1.	Balligand, J.-L., D. Ungureanu, R. A. Kelly, L. Kobzik, D. Pimental, T. Michel, and T. W. Smith. Abnormal contractile function due to induction of nitric oxide synthesis in rat cardiac myocytes follows exposure to activated macrophage-conditioned medium. J. Clin. Invest. 91: 2314-2319, 1993.
2.	Balligand, J.-L., D. Ungureanu, W. Simmons, D. Pimental, T. Malinski, M. Kapturczak, Z. Taha, J. Lowenstein, A. Davidoff, R. Kelly, T. Smith, and T. Michel. Cytokine-inducible nitric oxide synthase (iNOS) expression in cardiac myocytes. J. Biol Chem. 269: 27580-27588, 1994.
3.	Balon, T. W., and J. L. Nadler. Nitric oxide release is present from incubated skeletal muscle preparations. J. Appl. Physiol. 77: 2519-2521, 1994.
4.	Beckman, J. S., T. W. Beckman, J. Chen, P. A. Marshall, and B. A. Freeman. Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide. Proc. Natl. Acad. Sci. USA 87: 1620-1624, 1990.
5.	Boczkowski, J., B. Dureuil, C. Branger, D. Pavlovic, D. Murciano, R. Pariente, and M. Aubier. Effects of sepsis on diaphragmatic function in rats. Am. Rev. Respir. Dis. 138: 260-265, 1988.
6.	Bosenberg, A. T., J. G. Brock-Utne, S. L. Gaffin, M. T. B. Wells, and G. T. W. Blake. Strenuous exercise causes systemic endotoxemia. J. Appl. Physiol. 65: 106-108, 1988.
7.	Brady, A. J. B., P. A. Poole-Wilson, S. E. Harding, and J. B. Warren. Nitric oxide production within cardiac myocytes reduces their contractility in endotoxemia. Am. J. Physiol. 263 (Heart Circ. Physiol. 32): H1963-H1966, 1992.
8.	Brady, A. J. B., J. B. Warren, P. A. Poole-Wilson, T. J. Williams, and S. E. Harding. Nitric oxide attenuates cardiac myocyte contraction. Am. J. Physiol. 265 (Heart Circ. Physiol. 34): H176-H182, 1993.
9.	Buttery, L. D., T. Evans, D. Springall, A. Carpenter, J. Cohen, and J. M. Polak. Immunochemical localization of inducible nitric oxide synthase in endotoxin-treated rats. Lab. Invest. 71: 755-764, 1994.
10.	Cannon, J. G., R. A. Fielding, M. A. Fiatarone, S. F. Orencole, C. A. Dinarello, and W. E. Evans. Increased interleukin-1 in human skeletal muscle after exercise. Am. J. Physiol. 257: R451-R455, 1989.
11.	Chomczynski, P., and N. Sacchi. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162: 156-159, 1987.
12.	Cleeter, M. W. J., J. M. Cooper, V. M. Darley-Usmar, S. Moncada, and A. H. V. Shapira. Reversible inhibition of cytochrome-c oxidase, the terminal enzyme of the mitochondria respiratory chain by nitric oxide. FEBS Lett. 345: 50-54, 1994.
13.	Diaz, P. T., M. W. Julian, M. D. Wewers, and T. L. Clanton. Tumor necrosis factor and endotoxin do not directly affect in vitro diaphragm function. Am. Rev. Respir. Dis. 148: 281-287, 1993.
14.	Dufaux, B., and U. Order. Plasma elastase-1-antitrypsin, neopterin, tumor necrosis factor, and soluble interleukin-2 receptor after prolonged exercise. Int. J. Sports Med. 10: 434-438, 1989.
15.	Evans, H. G., M. J. Lewis, and A. M. Shah. Interleukin-1 modulates myocardial contraction via dexamethasone sensitive production of nitric oxide. Cardiovasc. Res. 27: 1486-1490, 1993.
16.	Finkel, M. S., C. V. Oddis, T. D. Jacob, S. C. Watkins, B. G. Hattler, and R. L. Simmons. Negative inotropic effects of cytokines on the heart mediated by nitric oxide. Science Wash. DC 257: 387-389, 1992.
17.	Freeman, B. Free radical chemistry of nitric oxide: looking at the dark side. Chest 105, Suppl.: 79S-84S, 1994.
18.	Geller, D. A., C. J. Lowenstein, R. A. Shapiro, A. K. Nussler, M. Di Silvio, S. C. Wang, D. K. Nakayama, R. L. Simmons, S. H. Snyder, and T. R. Billiar. Molecular cloning and expression of inducible nitric oxide synthase from human hepatocytes. Proc. Natl. Acad. Sci. USA 90: 3491-3495, 1993.
19.	Hauptmann, S., B. Klosterhalfen, J. Weis, C. Mittermayer, and C. J. Kirkpatrick. Skeletal muscle oedema and muscle fibre necrosis during septic shock. Observations with a porcine septic shock model. Virchows Archiv. 424: 653-659, 1994.
20.	Hussain, S. N. A., R. Graham, F. Rutledge, and C. Roussos. Respiratory muscle energetics during endotoxic shock in dogs. J. Appl. Physiol. 60: 486-493, 1986.
21.	Hussain, S. N. A., G. Simkus, and C. Roussos. Respiratory muscle fatigue: a cause of ventilatory failure in septic shock. J. Appl. Physiol. 58: 2033-2040, 1985.
22.	Kim, W. S., M. E. Ward, and S. N. A. Hussain. Pathological O2 supply dependence of diaphragmatic and systemic O2 uptake during endotoxemia. J. Appl. Physiol. 77: 1093-1100, 1994.
23.	Kinugawa, K., T. Takahashi, O. Kohmoto, A. Yao, T. Aoyagi, S. Momomura, Y. Hirata, and T. Serizawa. Nitric oxide-mediated effects of interleukin-6 on [Ca2+]i and cell contraction in cultured chick ventricular myocytes. Circ. Res. 75: 285-295, 1994.
24.	Kobzik, L., M. B. Reid, D. S. Bredt, and J. S. Stamler. Nitric oxide in skeletal muscle. Nature Lond. 372: 546-548, 1994.
25.	Kröncke, K.-D., K. Fehsel, T. Schmidt, F. T. Zenke, I. Dasting, J. R. Wesener, H. Bettermann, K. D. Breunig, and V. Kolb-Bachofen. Nitric oxide destroys zinc-sulfur clusters inducing zinc release from metallothionein and inhibition of the zinc finger-type yeast transcription activator LAC9. Biochem. Biophys. Res. Commun. 200: 1105-1110, 1994.
26.	Lowenstein, C. J., J. L. Dinerman, and S. H. Snyder. Nitric oxide: a physiologic messenger. Ann. Int. Med. 120: 227-237, 1994.
27.	Lyons, C. R., G. J. Orloff, and J. M. Cunningham. Molecular cloning and functional expression of an inducible nitric oxide synthase from a murine macrophage cell line. J. Biol. Chem. 267: 6370-6374, 1992.
28.	Moncada, S., R. M. J. Palmer, and E. A. Higgs. Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol. Rev. 43: 109-142, 1991.
29.	Nakane, M., H. H. Schmidt, J. S. Pollock, U. Forstermann, and F. Murad. Cloned human brain nitric oxide synthase is highly expressed in skeletal muscle. FEBS Lett. 316: 175-180, 1993.
30.	Nathan, C. Nitric oxide as a secretory product of mammalian cells. FASEB J. 6: 3051-3064, 1994.
31.	Nathan, C., and Q. Xie. Nitric oxide synthases: roles, tolls and controls. Cell 78: 915-918, 1994.
32.	Paulus, W. J., P. J. Vantrimpont, and A. M. Shah. Acute effects of nitric oxide on left ventricular relaxation and diastolic distensibility in humans: assessment by bicoronary sodium nitroprusside infusion. Circulation 89: 2070-2078, 1994.
33.	Reid, M. B., K. E. Haack, K. M. Franchek, P. A. Valberg, L. Kobzik, and M. S. West. Reactive oxygen in skeletal muscle. I. Intracellular oxidant kinetics and fatigue in vitro. J. Appl. Physiol. 73: 1797-1804, 1992.
34.	Reid, M. B., T. Shoji, M. R. Moody, and M. L. Entman. Reactive oxygen in skeletal muscle. II. Extracellular release of free radicals. J. Appl. Physiol. 73: 1805-1809, 1992.
35.	Robbins, R. A., P. J. Barnes, D. R. Springall, J. B. Warren, O. J. Kwon, L. D. K. Buttery, A. J. Wilson, D. A. Geller, and J. M. Polak. Expression of inducible nitric oxide in human lung epithelial cells. BBRC 203: 209-218, 1994.
36.	Salter, M., R. G. Knowles, and S. Moncada. Widespread tissue distribution, species distribution and changes in activity of Ca++ dependent and Ca++ independent nitric oxide synthases. FEBS Lett. 291: 145-149, 1991.
37.	Segal, S. S. Invited editorial on "Nitric oxide is present from incubated skeletal muscle preparations". J. Appl. Physiol. 77: 2517-2518, 1994.
38.	Snyder, S. H. Nitric oxide. More jobs for that molecule. Nature Lond. 372: 504-505, 1994.
39.	Weyrich, A. S., X. Ma, M. Buerke, T. Murohara, V. E. Armstead, A. M. Lefer, J. M. Nicolas, A. P. Thomas, D. J. Lefer, and J. Vinten-Johansen. Physiological concentrations of nitric oxide do not elicit an acute negative inotropic effect in unstimulated cardiac muscle. Circ. Res. 75: 692-700, 1994.
40.	Wilcox, P., S. Osborne, and B. Bressler. Monocyte inflammatory mediators impair in vitro hamster diaphragm contractility. Am. Rev. Respir. Dis. 146: 462-466, 1992.
41.	Williams, G., T. Brown, L. Becker, M. Prager, and B. P. Giroir. Cytokine-induced expression of nitric oxide synthase in C2C12 skeletal muscle myocytes. Am. J. Physiol. 267 (Regulatory Integrative Comp. Physiol. 36): R1020-R1025, 1994.