Endotoxin-induced skeletal muscle contractile dysfunction:
contribution of nitric oxide synthases
Q.
El-Dwairi1,
A.
Comtois2,
Y.
Guo1, and
S. N. A.
Hussain1
1 Critical Care and Respiratory
Divisions, Royal Victoria Hospital and Meakins-Christie Laboratories,
McGill University, Montreal, Quebec H3A 1A1; and
2 Respiratory Division, Notre-Dame
Hospital, Université de Montréal, Montreal, Quebec, Canada
H3C 3J7
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ABSTRACT |
The aims of this
study were to assess the role of nitric oxide (NO) and the contribution
of different NO synthase (NOS) isoforms in skeletal muscle contractile
dysfunction in septic shock. Four groups of conscious rats were
examined. Group 1 served as control; groups 2, 3, and
4 were injected with
Escherichia coli endotoxin [lipopolysaccharide (LPS), 20 mg/kg ip] and killed after 6, 12, and 24 h, respectively. Protein expression was assessed by
immunoblotting and immunostaining. LPS injection elicited a transient
expression of the inducible NOS isoform, which peaked 12 h after LPS
injection and disappeared within 24 h. This expression coincided with a significant increase in nitrotyrosine formation (peroxynitrite footprint). Muscle expression of the endothelial and neuronal NOS
isoforms, by comparison, rose significantly and remained higher than
control levels 24 h after LPS injection. In vitro measurement of muscle
contractility 24 h after LPS injection showed that incubation with NOS
inhibitor (S-methyliosothiourea)
restored the decline in submaximal force generation, whereas maximal
muscle force remained unaffected. We conclude that NO plays a
significant role in muscle contractile dysfunction in septic animals
and that increased NO production is due to induction of the inducible
NOS isoform and upregulation of constitutive NOS isoforms.
endotoxin; sepsis; shock
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INTRODUCTION |
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.
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METHODS |
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
NG-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
NG-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.
Nitrotyrosine formation was also quantified by slot blotting.
Diaphragmatic crude homogenates (100 µg) were mixed with an equal
volume of sample buffer, boiled for 5 min, and loaded on nitrocellulose
membrane using slot-blotting apparatus (Life Technologies, Gaithersburg, MD). Membranes were then blocked with 5% nonfat dry milk
and probed overnight with primary polyclonal antinitrotyrosine antibody
(Upstate Biotechnology; 1 µg/ml) and horseradish
peroxidase-conjugated anti-rabbit secondary antibodies. Negative
controls were produced by replacing antinitrotyrosine antibody with a
rabbit immunoglobulin G (IgG). The specificity of antinitrotyrosine
antibody was assessed by preincubation with 10 mM authentic
nitrotyrosine (Alexis, San Diego, CA). Specific proteins were detected
with the ECL kit, and the intensity of the slots was evaluated by
densitometry, as mentioned above.
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 |
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.

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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.
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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).

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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.
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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).

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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.
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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.
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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.

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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.
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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).

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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.
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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.
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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).

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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.
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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.
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DISCUSSION |
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.
Using L-citrulline assay as an
indicator of iNOS activity, Salter et al. (29) reported for the first
time that diaphragmatic iNOS activity rose significantly 6 h after
inoculation of rats with Salmonella
typhimurium endotoxin. In a more recent
study, Boczkowski et al. (5) described iNOS expression in the diaphragm of rats injected with E. coli LPS.
Interestingly, early iNOS expression (within 6 h of LPS injection)
localized mainly in inflammatory cells infiltrating the diaphragm,
whereas myofibrillar iNOS expression was evident after 12 and 24 h (5).
iNOS expression was not detected in limb or ventilatory muscles other
than the diaphragm. We observed transient iNOS expression in the
diaphragm, which peaked 12 h after LPS injection and disappeared within
24 h. The expression of iNOS protein on Western blots was associated
with a significant rise in
Ca2+/calmodulin-independent NOS
activity (Fig. 1). Not only was the time course of iNOS expression in
the diaphragm in our study different from that in the study of
Boczkowski et al., but we detected significant iNOS induction in the
intercostal and limb muscles, whereas only diaphragmatic iNOS
expression was observed by Boczkowski et al. We speculate that these
differences between the two studies are related to variations in the
dose and serotype of E. coli
endotoxin, which could elicit different time courses of inflammatory
cytokine production. The two studies, however, confirm that
myofibrillar iNOS expression lags behind that of inflammatory cells
infiltrating muscle fibers. Similarly, Thompson et al. (34) failed to
detect iNOS expression in skeletal muscle fibers, despite abundant iNOS protein in the endothelium and macrophages of mice injected with E. coli LPS. The molecular mechanisms
responsible for iNOS induction in muscle fibers have not been addressed
in our study. We speculate, however, that activation of the iNOS
promoter in skeletal muscle fibers is mediated through mobilization of
the transcription factor nuclear factor-
B through pathways involving
tyrosine kinases and protein kinase C. The importance of these pathways
in iNOS induction in L6 cultured myoblasts was confirmed recently (27).
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).
To the best of our knowledge, our results provide the first evidence of
nitrotyrosine formation in normal and septic skeletal muscles. It is
interesting that the time course of increased nitrotyrosine formation
in the diaphragm and intercostal muscles was similar to that of iNOS
expression, suggesting that iNOS was the isoform responsible for
peroxynitrite formation in our experiments. On the other hand, we
speculate that the low level of nitration in the controls was due to
the activity of constitutively expressed NOS isoforms. Our results also
indicate that enhanced nitration of tyrosine residues in septic muscles
is a selective process and involves the nitration of two major proteins
with apparent molecular masses of ~196 and 86 kDa. The
nature of these proteins remains to be elucidated.
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).
Our data indicate that LPS injection elicited a substantial rise in
muscle NOS activity, which was due mainly to augmented Ca2+/calmodulin-dependent NOS
activity (Fig. 1). We attribute this finding to increased protein
expression of the ecNOS and nNOS isoforms as detected by immunoblotting
with isoform-selective antibodies. The heightened expression of these
isoforms was not limited to the diaphragm, since similar changes in
nNOS and ecNOS expression were also seen in other muscles such as the
intercostals, soleus, and gastrocnemius. Little is known about the
influence of LPS and inflammatory cytokines on the regulation of
skeletal muscle nNOS and ecNOS isoforms. In endothelial cells,
cytokines and LPS increase NO production, presumably via enhanced
activation and/or expression of the ecNOS isoform (17, 23).
Upregulation of ecNOS transcription rate in response to LPS and
cytokines has been confirmed recently in bovine aortic endothelial
cells (16). Similarly, there is evidence that in vivo exposure to LPS
(24) or in vitro administration of interferon-
(1) leads to
upregulation of nNOS expression in the brain. Several mechanisms are
likely to be involved in the upregulation of muscle nNOS and ecNOS
protein expression in septic animals. Our study indicates that one of these mechanisms is increased mRNA concentration of these isoforms (Fig. 5). More research is needed to elucidate whether the increase in
mRNA expression of these isoforms is due to improved mRNA stability and/or increased transcription rate. In addition, we need to
investigate the exact transcription factors involved in the regulation
of nNOS and ecNOS gene expression in response to endotoxemia or
inflammatory cytokines.
It should be emphasized that, unlike the diaphragm, intercostal, and
gastrocnemius muscles, total NOS activity of the soleus muscle did not
increase after LPS injection (Fig. 1). The reasons behind this
observation are unclear. We speculate that we were unable to detect a
significant rise in soleus total NOS activity after LPS injection
because this muscle expresses very low levels of constitutive NOS
isoforms, especially nNOS. Indeed, in the control group of animals, NOS
activity was lower in the soleus than in the muscles (Fig. 1). Similar
findings have been reported by Kobzik et al. (19). Moreover, we
recently compared nNOS expression in rat muscles and observed very low
nNOS expression in the soleus compared with the intercostal, diaphragm,
and gastrocnemius muscles (15). Thus it is possible that LPS-induced
changes in soleus constitutive NOS expression might not have been large
enough to be detected with the
L-citrulline assay. It is also
likely that variations in local cytokine levels, the degree of
inflammatory cell infiltration, and other unknown factors may have
contributed to the lack of a significant rise in soleus NOS activity
after LPS injection.
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.
The exact sites through which NO influences muscle force remains to be
elucidated. There is evidence, however, that NO regulates muscle
contractile force through guanosine 3',5'-cyclic
monophosphate (cGMP)-dependent and cGMP-independent processes, which
involve Ca2+ release by ryanodine
receptors (25). This notion is supported by the finding that agents
that increase intracellular cGMP, such as 8-BrcGMP, reverse the
influence of NOS inhibitors on muscle force (19). It has also been
proposed that NO depresses mitochondrial respiration in skeletal
muscles by inhibiting cytochrome c
oxidase, a process that is likely to decrease cellular ATP and increase ADP, AMP, GDP, and Pi. These
metabolites regulate diverse cellular processes, such as ion transport,
protein synthesis, and muscle contraction. NO could also modulate
muscle contractility by inhibiting creatine kinase (39). Finally, it
has been proposed that peroxynitrite, formed by NO and superoxide
anions, is involved in sepsis-induced skeletal muscle dysfunction
through increased lipid peroxidation and oxygen free radical damage (5,
31). Our results confirm that endotoxemia leads to enhanced tyrosine
nitration of the ventilatory muscles of septic rats, which coincided
with iNOS expression in these muscles.
The source of enhanced NO production in endotoxemic muscles has not
been identified. Boczkowski et al. (5) and Gath et al. (11) attributed
LPS-induced muscle contractile dysfunction to iNOS induction in muscle
fibers. Although our results confirm iNOS induction in skeletal
muscles, upregulation of nNOS and ecNOS protein expressions and the
significant rise in
Ca2+/calmodulin-dependent NOS
activity in these muscles suggest that iNOS may not be the only source
of enhanced NO production in septic animals. This is particularly true
in the late phase of septic shock (24 h), where iNOS expression could
no longer be detected and ecNOS and nNOS expression peaked. The
functional significance of this rise in ecNOS and nNOS expression
remains to be assessed by using selective gene knockouts of NOS
isoforms or potent isoform-selective NOS inhibitors.
In summary, our study indicates that LPS-induced muscle contractile
dysfunction is due to enhanced NO release, which, in turn, could be
attributed to iNOS induction and upregulation of ecNOS and nNOS
expressions. We also found that iNOS induction in muscle tissues was
associated with nitrotyrosine formation, suggesting that iNOS induction
is involved in muscle peroxynitrite production.
 |
ACKNOWLEDGEMENTS |
We are grateful to J. Longo, R. Carin, and O. DaSilva for help in
editing the manuscript.
 |
FOOTNOTES |
This study was funded by a grant from the Medical Research Council of
Canada. S. N. A. Hussain is a scholar of Fonds de la Recherche au
Santé du Quebec.
Address for reprint requests: S. N. A. Hussain, Rm. L3.05, Royal
Victoria Hospital, 687 Pine Ave. West, Montreal, PQ, Canada H3A 1A1.
Received 19 August 1997; accepted in final form 11 December 1997.
 |
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