Approximately half of the nNOSμ protein expressed in skeletal muscle is bound to the sarcolemmal dystrophin-glycoprotein complex, while the reminder resides in the cytoplasm (1
). To determine the location of cytoplasmic nNOS, we immunolabeled single muscle fibers (Figure A), using a method optimized for the resolution of subcellular organelles (23
). Four pan-specific anti-nNOS antibodies raised against distinct epitopes detected nNOS localized to sub-sarcolemmal puncta (Figure , B and C, and Supplemental Figure 1; supplemental material available online with this article; doi:
). Sarcolemmal nNOSμ labeling is present in this fiber but is out of the plane of focus. The punctate labeling pattern resembled that of the murine skeletal muscle Golgi complex (23
). In fact, nNOS puncta colocalized precisely with GM130, a marker of the cis
-face of the Golgi complex in fast-twitch gastrocnemius (Figure B) and slow-twitch soleus muscle fibers (Figure C). These data suggest the existence of a previously unrecognized Golgi-associated nNOS compartment in skeletal muscle cells.
nNOS is localized to the Golgi complex in fast- and slow-twitch skeletal muscle fibers.
To establish the identity of the Golgi nNOS, we used nNOS1 knockout 1 (KN1, knockout of nNOSμ only; Figure D) and nNOS knockout 2 (KN2, knockout of any active full-length nNOS splice variants; Figure E) mice (see Methods and refs. 24
). At least 4 splice variants of nNOS may be transcribed from the Nos1
gene; however, nNOSα is not expressed in skeletal muscle, leaving only nNOSμ, nNOSβ, and nNOSγ as possible candidates for Golgi nNOS (Figure A). Surprisingly, Golgi nNOS labeling was unaffected in KN1 skeletal muscles (Figure A), indicating that the Golgi nNOS was not nNOSμ. Golgi nNOS labeling was absent in skeletal muscles from KN2 mice, further confirming nNOS antibody specificity (Figure A). Taken together, these data suggest that the Golgi-associated nNOS is either nNOSβ and/or nNOSγ. While we cannot definitively eliminate nNOSγ as a candidate for the Golgi nNOS, it seems unlikely. Unlike nNOSβ, which is catalytically normal and possesses a unique amino terminus, nNOSγ is enzymatically inactive and lacks unique primary sequence to facilitate Golgi targeting (Figure A) (4
). Therefore, throughout this paper, we will refer to skeletal muscle Golgi nNOS as nNOSβ. More importantly, these data suggest the existence of 3 distinct nNOS compartments (Golgi nNOSβ, sarcolemmal nNOSμ, and cytoplasmic nNOSμ) and reveal the previously unrecognized complexity of nNOS-based signaling in skeletal muscle.
nNOS splice variants and nNOS mutant mouse lines used to analyze NO signaling in skeletal muscle.
A nNOS splice variant localizes to the Golgi complex and regulates microtubule cytoskeleton integrity in skeletal muscle.
Classical NO signal transduction is mediated by NO-sensitive sGC and PKG. To test the possibility that NO-cGMP signaling could occur at the skeletal muscle Golgi complex, we examined whether sGC and PKG localized to Golgi membranes. Both sGC (Figure A) and PKG (Figure B) colocalized with GM130. These data suggest that sGC and PKG, the primary targets and effectors of NO, are in close proximity to nNOSβ at the Golgi. These data strongly support the possible existence of a novel nNOS-cGMP signal transduction microdomain at the Golgi complex in skeletal muscle.
A NO-cGMP signaling microdomain that we believe to be novel at the Golgi complex in skeletal muscle.
Skeletal muscles from KN1 and KN2 mice were examined to determine the subcellular consequences of nNOS splice variant deficiency on skeletal muscle organization. The absence of nNOS in the muscles of KN2 mice disrupted the distribution of the Golgi (Figure A, bottom row), which was unaffected in KN1 relative to WT muscle (Figure A, middle and top rows). Since microtubules are responsible for Golgi complex localization (26
), we evaluated microtubule cytoskeleton organization in nNOS mutant mice (Figure B). The orthogonal lattice of microtubules was unaffected by the absence of nNOSμ (Figure B). In contrast, the microtubule cytoskeleton was dramatically disrupted in nNOS-deficient muscles of KN2 mice (Figure B). These data suggest that nNOSβ regulates microtubule cytoskeleton integrity. nNOSβ deficiency contributes to destabilization of the microtubule lattice and perturbation of Golgi membrane organization.
Muscle fatigue resistance is an excellent global measure of muscle performance. We evaluated this parameter in tibialis anterior (TA) muscles in situ from KN1, KN2, and α-syntrophin–null mice (Figure C and Figure and Table ). nNOSμ fails to localize to the sarcolemma in α-syntrophin–null muscle, despite preservation of nNOSμ expression and activity; therefore α-syntrophin knockouts are especially useful in specifically figuring out sarcolemmal nNOSμ function (7
). Given the importance of contraction-induced nNOSμ signaling in vascular function, we used an in situ method for testing muscle contractile function, in which normal vascularization and innervation of skeletal muscle tissue are maintained. After 4 minutes of simulated exercise, force output from WT TA muscles was 50% of initial levels (Figure and Table ). The loss of sarcolemmal nNOSμ only in the α-syntrophin knockout (Figure C) had no significant impact on contraction-induced fatigue or postexercise force generation despite aberrant vasoconstriction (Figure and Table ) (7
). Contrary to expectation, the absence of muscle nNOSμ in KN1 mice also had no impact on muscle fatigue or postexercise strength (Figure and Table ). Using an identical approach, we previously reported mild contraction-induced fatigue in nNOSμ-deficient TA muscles in KN1 mice on a B6129 background (10
). The mice in this study were all generated on, or backcrossed 10 generations onto, a C57BL/6 background. These data demonstrate strain-specific modulation of nNOSμ-regulated fatigue. Our data also argue that the functional ischemia in the absence of contraction-induced sarcolemmal nNOSμ signaling does not affect susceptibility to contraction-induced fatigue or postexercise force generating capacity.
nNOS splice variants differentially regulate contraction-induced fatigue and postexercise force recovery.
Skeletal muscle fatigue in KN1, KN2, and α-syntrophin–null mice
In contrast, the loss of all nNOS splice variants in KN2 muscle dramatically increased susceptibility to contraction-induced fatigue (Figure and Table ). KN2 skeletal muscles also exhibited significant force deficits after exercise. Postexercise weakness was apparent in KN2 muscles even at 5 minutes after exercise (Figure and Table ). These data provide 4 insights, which we believe to be new, into nNOS splice variant function: (a) loss of contraction-induced signaling from nNOSμ does not necessarily limit force production during or after exercise; (b) nNOSβ is a critical regulator of skeletal muscle fatigue and postexercise force output; (c) during exercise, nNOSμ signaling maintains blood delivery to active muscle, while nNOSβ regulates muscle fatigue resistance and postexercise force output; and (d) the differential targeting of nNOS splice variants creates functionally distinct NO signaling microdomains, at which NOS-derived NO acts locally in skeletal muscle.
A possible molecular explanation for increased muscle fatigue in KN2 muscle is a decrease in the ratio of fatigue-resistant to fatigue-susceptible muscle fibers. To address the possibility that altered fiber composition contributes to increased KN2 muscle fatigue, we determined the frequency of fast- and slow-twitch fibers in nNOS mutant TA muscles (Figure A). Fibers expressing type I myosin heavy chain (MyHC) are the most fatigue resistant, while those expressing type IIb MyHC are the least fatigue resistant (28
). Type IIa–positive myofibers exhibit an intermediate fatigue resistance. Type IIx/IId fibers are also relatively resistant to fatigue (28
). KN1 muscles exhibited a shift to a more fatigue-resistant fiber composition, characterized by a significant 50% reduction in type IIa–positive fibers and a remarkable 3,000% increase in type IIx/IId fibers compared with normal controls (Figure B and Supplemental Figure 2). The marked increase in type IIx/IId fibers could contribute to the fatigue resistance of nNOSμ-deficient KN1 TA muscle. Faster mean relaxation and peak twitch force generation times are also consistent with the increased type IIx/IId fast fiber composition of KN1 TA muscle (Supplemental Figure 3). In contrast, there was a significant 45% increase relative to WT in type IIb fibers in nNOS-deficient KN2 muscles that could contribute to decreased fatigue resistance. The shift toward more fatigue-resistant and fatigue-susceptible fiber compositions in KN1 and KN2 muscles, respectively, provides a potential mechanism for the differential impact of nNOS splice variants on contraction-induced muscle fatigue.
nNOS-deficient KN2 skeletal muscles exhibit a more fatigue-susceptible fiber composition.
In order to determine the full extent of nNOS splice variant regulation of skeletal muscle contractile function, we determined maximal tetanic force generating capacity and specific force (total tetanic force normalized to muscle cross-sectional area [CSA]) for all nNOS genotypes. Maximum tetanic force output did not differ significantly between WT, α-syntrophin–deficient and KN1 TA muscles (Figure A). In contrast, KN2 muscles lacking nNOSμ and nNOSβ generated only 40% of the maximum isometric force of WT controls (Figure A). Similarly, KN2 muscles were significantly weaker, exhibiting a 14% reduction in mean specific force (Figure B). Thus, nNOSβ-derived NO appears necessary to maintain normal skeletal muscle strength.
Golgi nNOS splice variant is required to maintain normal skeletal muscle strength, muscle mass, and myofiber size.
Since muscle strength correlates positively with muscle size, we investigated whether the intrinsic weakness of KN2 muscle was due to a reduction in muscle mass. The mass of α-syntrophin–null and KN1 TA muscles did not differ from controls (Figure C); however KN2 muscle mass decreased 40% (P < 0.001) compared with WT controls. Normalization of muscle mass to body weight demonstrated that the decrease was not simply attributable to decreased body mass (data not shown). The median CSA of KN2 TA myofibers was significantly smaller (1,382 μm2) than that of controls (1,773 μm2) and KN1 muscle cells (1,814 μm2; Figure D). Thus, decreased muscle mass in KN2 mice is likely due to a reduction in the size of the muscle cells themselves.
We then tested whether all fiber types were equally reduced in size by the absence of nNOSβ (Supplemental Figure 2). We measured the CSA of type IIa and type IIb fibers, which together make up approximately 90% of the myofibers in WT and KN2 TA muscles. The CSA of type IIb fibers was significantly decreased by 31% (P < 0.05) in KN2 TA muscles, while the area of type IIa fibers was not significantly affected (Supplemental Figure 2B). It is important to note the 45% increase in the number of type IIb fibers per square millimeter in KN2 TA muscles (Figure and Supplemental Figure 2); therefore, the decrease in KN2 TA muscle mass is not simply due to a change in fiber type but is attributable to, at least in part, increased numbers of significantly smaller type IIb fibers. In contrast, nNOSμ deficiency resulted in a significant decrease in KN1 soleus muscle mass and median CSA relative to WT (Supplemental Figure 4). However, similar to the TA, KN2 soleus muscles were also smaller in mass and CSA (Supplemental Figure 4). These data suggest that nNOSβ signaling pathways regulate muscle mass and myofiber size independently of muscle type, whereas nNOSμ signaling does so through muscle type-specific mechanisms.
Destabilization of the microtubule cytoskeleton, mislocalization of the Golgi complex, and reduced myofiber size were obvious myopathic changes evident from light microscopy-based analyses of WT, KN1, and KN2 muscles (Figure A and Supplemental Figure 4). To identify additional myopathic changes in muscle cytoarchitecture, we examined skeletal muscles at the ultrastructural level. Electron microscopic analysis of KN1 TA muscle revealed intermyofibrillar mitochondria that were swollen and electron lucent (presumably reflecting decreased matrix density) (Figure B). Sarcomere organization and registration in KN1 muscle were indistinguishable from that of controls (Figure B). In KN2 TA muscles, intermyofibrillar mitochondria were often more swollen and variable in size, electron lucent, and aberrantly localized (Figure B). This is consistent with defects in the organization of the microtubule cytoskeleton that anchors mitochondrial organelles (Figure B) (29
). Furthermore, regions of muscle appeared highly disorganized, with impaired sarcomere alignment. To test whether structural abnormalities in nNOS-deficient mice lead to increased muscle fragility, thus predisposing TA muscles to contraction-induced injury, force output was determined at progressively increasing strains. α-Syntrophin–null, KN1, and KN2 muscles exhibited force deficits that were indistinguishable from that of WT controls (Supplemental Figure 5). Similarly, creatine kinase activity was unaffected in the serum from KN1 (ref. 18
and data not shown) and KN2 (Supplemental Figure 6) mice, indicating normal myofiber stability and turnover. Together, these data show that nNOS splice variant deficiency does not increase susceptibility to contraction-induced injury or decrease muscle cell stability, in agreement with a previous study (10
), and do suggest an important role for both nNOSμ and nNOSβ splice variants in maintaining normal mitochondrial health.
nNOS splice variant-deficiency leads to myopathic changes in intermyofibrillar mitochondria and skeletal muscle cytoarchitecture.