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Journal of Applied Physiology
Vol. 82, No. 4, pp. 1250-1255, April 1997
EXERCISE AND MUSCLE

Induction of neuronal type nitric oxide synthase in skeletal muscle by chronic electrical stimulation in vivo

Peter J. Reiser1, William O. Kline2, and Pal L. Vaghy3

1 Department of Oral Biology, 2 Department of Exercise Science, and 3 Department of Medical Biochemistry, The Ohio State University, Columbus, Ohio 43210-1218

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
ACKNOWLEDGEMENTS
FOOTNOTES
REFERENCES

ABSTRACT

Reiser, Peter J., William O. Kline, and Pal L. Vaghy. Induction of neuronal type nitric oxide synthase in skeletal muscle by chronic electrical stimulation in vivo. J. Appl. Physiol. 82(4): 1250-1255, 1997.---Fast-twitch skeletal muscles contain more neuronal-type nitric oxide synthase (nNOS) than slow-twitch muscles because nNOS is present only in fast (type II) muscle fibers. Chronic in vivo electrical stimulation of tibialis anterior and extensor digitorum longus muscles of rabbits was used as a method of inducing fast-to-slow fiber type transformation. We have studied whether an increase in muscle contractile activity induced by electrical stimulation alters nNOS expression, and if so, whether the nNOS expression decreases to the levels present in slow muscles. Changes in the expression of myosin heavy chain isoforms and maximum velocity of shortening of skinned fibers indicated characteristic fast-to-slow fiber type transformation after 3 wk of stimulation. At the same time, activity of NOS doubled in the stimulated muscles, and this correlated with an increase in the expression of nNOS shown by immunoblot analysis. These data suggest that nNOS expression in skeletal muscle is regulated by muscle activity and that this regulation does not necessarily follow the fast-twitch and slow-twitch pattern during the dynamic phase of phenotype transformation.

nitric oxide; enzyme induction; striated muscles; membranes

INTRODUCTION

NEURONAL-TYPE NITRIC OXIDE SYNTHASE (nNOS) is present in all striated muscles of mammals (11). Some skeletal muscles express more nNOS than any other tissue, including brain (19). Because skeletal muscle represents ~40% of the total body mass (12), this tissue may be one of the richest sources of nNOS and nitric oxide (NO) in mammals. The nNOS is associated with the sarcolemma of fast (type II) skeletal muscle fibers (15). Consistent with this finding is that expression and activity of nNOS are high in muscles rich in type II fibers (fast muscles) and low in muscles rich in type I fibers (slow muscles) (6, 15). In addition to nNOS, skeletal muscle also contains endothelial nitric oxide synthase (eNOS), albeit at much lower levels (16). The nNOS is associated with the dystrophin complex by binding to alpha 1-syntrophin in fast-twitch muscle fibers (5, 6). In mdx mice, an animal model of muscular dystrophy, and in humans with Duchenne muscular dystrophy, where the expression of dystrophin is altered or missing, the nNOS dissociates from the sarcolemma (5, 6). It has been suggested that such displacement from the sarcolemma may result in an aberrant regulation of nNOS and muscle degeneration in Duchenne muscular dystrophy (6). Examination of nNOS under pathological conditions may lead to a better understanding of the role of this enzyme in skeletal muscle.

Although originally thought of as a constitutive enzyme, recent data indicate that expression and activity of nNOS are altered under different developmental and hormonal conditions (20, 30). Whether nNOS expression in type II skeletal muscle fibers can be changed by altering the muscle activity has not been determined. Chronic electrical stimulation of fast skeletal muscles is a strong enough stimulus to produce fast-to-slow fiber type transformation. We tested whether such electrical stimulation of fast skeletal muscles is able to alter the expression and activity of nNOS. The data suggest that chronic electrical stimulation of tibialis anterior (TA) and extensor digitorum longus (EDL) muscles increases the activity of nitric oxide synthase (NOS) as well as the level of nNOS expression. A brief report of this study was presented at the Second International Conference on Biochemistry and Molecular Biology of Nitric Oxide, Los Angeles, California, July 13-17, 1996.

MATERIALS AND METHODS

Stimulation protocol. Three adult female New Zealand White rabbits, weighing 2.5-3.5 kg, were used in this study. The rabbits were anesthetized with 60 mg/kg ketamine and 5 mg/kg xylazine before surgical implantation of electrodes. Continuous stimulation at 10 Hz with 0.1-ms biphasic pulses of the right peroneal nerve, which innervates the fast-twitch EDL and TA muscles, was started 3 days later and lasted 3 wk (2). The contralateral muscles were used as controls. The animals were killed at the end of the stimulation period with pentobarbital sodium (150 mg/kg iv), and the muscles were removed and processed as described below. The care and use of animals in this study were approved by the institutional laboratory animal care and use committee.

Myosin heavy chain (MHC) analysis. A portion (30-50 mg) of each muscle was prepared as previously described (4) up to the centrifugation step, except that the samples were homogenized after sample buffer was added. An aliquot of the supernatant was diluted 1:10 with sample buffer and 4 µl were loaded onto a sodium dodecyl sulfate-polyacrylamide gel for analysis of MHC isoform composition.

Electrophoresis was performed according to Talmadge and Roy (28) with the following modifications. The stacking gel included 5% (vol/vol) glycerol, and the upper electrode buffer included 10 mM 2-mercaptoethanol (10). The gels were run in a Hoefer Scientific SE 600 unit (Hoefer Scientific Instruments, San Francisco, CA) at 8°C for 24 h with constant voltage (275 V).

The gels were silver stained and dried as in Blough et al. (4) and analyzed by using a scanning densitometer (model GS 300, Hoefer Scientific Instruments) to quantify the relative amounts of MHC isoforms in the samples. The linearity of densitometric scanning of myosin heavy chain isoforms was tested by scanning gels on which varying loads of adductor magnus and diaphragm samples, prepared as described above, were electrophoresed. Adductor magnus and diaphragm muscles are composed of myosin heavy chain isoforms types IIB and IID, and types I, IIA, and IID, respectively (1). Linear correlation coefficients of 0.97 or greater were obtained for each MHC isoform when the densitometric areas were regressed against final (i.e., diluted) sample volumes ranging from 0.5 to 5 µl.

Immunoblot. nNOS expression was evaluated with modifications of the immunoblotting procedure of Towbin et al. (29). The separating and stacking gels were identical to those used in the MHC analysis, except that glycerol was not included in either gel. Equal total protein amounts (16 µg) of total membrane fractions were dissolved in the sample buffer and electrophoresed in a minigel system (Hoefer SE250). The proteins were then transferred to nitrocellulose membrane in a mini-V BRL unit (Life Technologies, Gaithersburg, MD). Blocking of the membranes was performed with 5% bovine serum albumin in 25 mM tris(hydroxymethyl)aminomethane (Tris), 192 mM glycine, 0.01% (wt/vol) sodium dodecyl sulfate, 15% (vol/vol) methanol, and 0.02% (vol/vol) Tween 20 [in Tris-buffered saline (TBST)] for 1 h. The membranes were reacted with anti-nNOS and anti-eNOS primary antibodies (N31030 and N30030, respectively; Transduction Laboratories, Lexington, KY) diluted in TBST. Membranes were washed three times, 5 min each, in TBST. Membranes were reacted with an alkaline phosphatase-conjugated secondary antibody (111-055-003, Jackson ImmunoResearch Laboratories, West Grove, PA) diluted 1:7,000 in TBST. Nitro-blue tetrazolium chloride (132 µl) and 66 µl 5-bromo-4-chloro-3-inodyl-1-phosphate in 20 ml of 0.1 M MgCl2, 1.5 M NaCl, and 1 M Tris were utilized for color development. All of the steps after completion of the transfer were performed with shaking at 37°C.

Maximal velocity of shortening (Vmax ) of single fibers. Bundles of fibers were prepared from the control and stimulated TA and EDL muscles, as well as from one soleus muscle, from one rabbit and stored in glycerinating solution at -22°C (23) to render the sarcolemma hyperpermeable. The portions were dissected from the same region of the control and stimulated muscles. Single skinned fibers were dissected from the bundles, and Vmax was measured by utilizing the slack-test method of Edman (9). Only those fibers that did not have morphological irregularities, when viewed under a microscope at ×40 magnification, were selected for measurements.

Membrane isolation. A sample of each muscle was weighed, minced, and diluted 10 times with isolation buffer [(in mM) 50 Tris · HCl, pH 7.4 at 4°C, 1 EDTA, 0.5 phenylmethylsulfonyl fluoride, and 1 iodoacetamide, as well as 1 µM leupeptin and 1 µM pepstatin A]. The minced tissue was homogenized with a polytron twice for 15 s at a setting of eight by using a PTA 10S generator. In between each 15-s homogenization, the tissue was kept at melting-ice temperature for 60 s. The homogenate was filtered through one layer of cheesecloth into an ultracentrifuge tube and centrifuged at 105,000 g for 1 h. The pellet was resuspended in isolation buffer.

NOS activity measurement. The citrulline assay (13), which directly measures one of the NOS products (citrulline) and can be performed with crude membrane preparations without the interference of other cellular components, was used. The assay was performed in 1.5-ml polypropylene bullet tubes incubated in a 37°C water bath. The final reaction mixture (120 µl) contained 55 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES)-NaOH, 20 mM Tris · HCl, pH 7.4, various concentrations of L-[14C(U)]arginine monohydrochloride (Amersham, Arlington Heights, IL), 25 µl skeletal muscle membrane, 0.8 mM 1,4-dithiothreitol, 0.4 mM EDTA, cofactors [0.8 µM NADPH, 1 mM CaCl2, 0.5 µM bovine brain calmodulin, 6 µM sapropterin (BH4), 0.8 µM flavin-adenine dinucleotide, 0.8 µM riboflavin monophosphate], and protease inhibitors (0.2 mM phenylmethylsulfonyl fluoride, 0.4 mM iodoacetamide, 0.4 µM leupeptin, 0.4 µM pepstatin A). The reaction was initiated by the addition of protein. The samples were incubated at 37°C for 20 min, unless otherwise indicated. The reaction was stopped by the addition of 1 ml of ice-cold stop solution (20 mM HEPES, pH 6.8, containing 2 mM EDTA). The citrulline produced by NOS was separated by cation-exchange chromatography by using 2-ml AG 50W-X8 columns (Bio-Rad) converted to the Na+ form and preequilibrated with the stop solution. The radioactivity was counted in a Beckman LS7000 liquid scintillation counter at 95% efficiency. The exact concentration of L-[14C(U)]arginine used in the assays was determined by measuring radioactivity in 20-ml aliquots taken from the reaction mixture not used in the assays. Control experiments included measurement of the NOS activity in the presence of stop solution and/or stereoselective NOS inhibitors such as Nomega -monomethyl-L-arginine (L-NMMA). The same control (blank) values were obtained if the membranes were incubated with stop solution, if the membranes were incubated with 500 µM L-NMMA, or if membranes were omitted from the samples. The protein concentration was determined by using the bicinchoninic acid protein assay reagent (Pierce, Rockford, IL).

Statistical analysis. Differences in MHC isoform composition, Vmax, and muscle mass were evaluated statistically with the one-tailed Mann-Whitney-Wilcoxon test after an analysis of variance. The level of significance was set at P < 0.05.

RESULTS

The mean mass of electrically stimulated TA and EDL muscles decreased by 40 ± 3 and 30 ± 7%, respectively, after 3 wk of electrical stimulation in vivo (Table 1). To determine whether fast-to-slow fiber type transformation took place, the expression of MHC isoforms and Vmax of isolated skinned fibers were analyzed. MHC isoforms were identified on the basis of their electrophoretic mobilities and relative abundance in control soleus, diaphragm, and adductor magnus muscles (1). Four MHC isoforms, IIA, IID, IIB, and I, were identified in order of their electrophoretic mobilities (Fig. 1A). The soleus expressed primarily MHC type I isoform. The diaphragm contained type IIA, type IID, and type I MHC isoforms. The adductor magnus primarily expressed the type IIB isoform. The four different MHC isoforms were best shown by coelectrophoresis of diaphragm and adductor magnus muscle samples (Fig. 1A).

Table 1. Stimulation-induced changes in muscle mass, myosin heavy chain isoforms, and shortening velocity in tibialis anterior and extensor digitorum longus muscles

Control TA Stimulated TA Control EDL Stimulated EDL Control Soleus
Muscle mass, g 3.28 ± 0.11  1.94 ± 0.07* 2.85 ± 0.18  2.01 ± 0.15* 2.16 ± 0.13 
%MHC I 5 ± 2  19 ± 5* 5 ± 1  27 ± 6* 84 ± 5 
%MHC IIA 39 ± 6  65 ± 7* 35 ± 3  55 ± 4* 16 ± 5 
%MHC IID 56 ± 7  15 ± 3* 60 ± 3  17 ± 3* 0
Single-fiber Vmax, fl/s 3.74 ± 0.29  2.42 ± 0.37* 4.17 ± 0.42  1.78 ± 0.36* 0.91 ± 0.02
Values are means ± SE; n = 3 except for maximal velocity of shortening [Vmax; n = 10 for both control and stimulated tibialis anterior (TA) muscles; n = 6 for control and stimulated extensor digitorum longus (EDL) muscles and control soleus muscle, respectively]. Values for myosin heavy chain (MHC) isoforms are expressed as %total MHC in each muscle. Vmax is expressed in fiber length (fl)/s. * Significantly different from control, P < 0.05.

Fig. 1. Myosin heavy chain (MHC) expression in control and stimulated rabbit skeletal muscles. A: relative amounts of MHC isoforms (IIA, IID, IIB, and I) in soleus (Sol), diaphragm (Dia), adductor magnus (AM), or a combination of adductor magnus and diaphragm (AM + Dia) muscles. B: MHC isoforms in nonstimulated, control (C) and stimulated (S) tibialis anterior (TA) and extensor digitorum longus (EDL) muscles.
[View Larger Version of this Image (37K GIF file)]

The MHC isoforms present in control and stimulated TA and EDL muscles are shown in Fig. 1B. Control TA and EDL muscles contained predominantly MHC types IIA and IID and relatively small amounts of type I MHC. The expression of type IID MHC was reduced and the expressions of type I and IIA MHC were increased in both muscles on chronic electrical stimulation (Fig. 1B). The percentages of total MHC corresponding to type I, IIA, and IID isoforms in control and stimulated TA and EDL muscles are shown in Table 1.

The mean Vmax of stimulated TA and EDL muscle fibers was significantly lower than that of the respective control muscles (Table 1). However, the Vmax of both stimulated TA and EDL muscle fibers was still greater after 3 wk of stimulation than that of soleus fibers (Table 1).

To validate the usefulness of the citrulline assay for quantitation of NOS activity of rabbit skeletal muscle fractions, the NOS activity was first measured in skeletal muscle membranes isolated from mixed rabbit leg and back muscles. Under our assay conditions, the rate of NOS activity increased linearly during the first 20 min of incubation (Fig. 2A). The NOS activity also increased linearly with the protein concentration up to at least 4 mg protein/ml sample (Fig. 2B). In agreement with the findings of others (15, 19), almost all NOS activity was associated with the membrane fraction (Fig. 2C). The supernatant obtained after sedimentation of insoluble materials by centrifugation at 105,000 g for 60 min contained <15% of the total activity present in rabbit skeletal muscle (Fig. 2C).

Fig. 2. Characterization of rabbit skeletal muscle nitric oxide synthase (NOS). A: time dependence of NOS activity. Skeletal muscle membranes (0.47 mg protein) were incubated with 3.26 mM L-[14C(U)]arginine at 37°C for various periods of time before assay. B: protein dependence of NOS activity. Various concentrations of rabbit skeletal muscle membranes were incubated with 3.27 mM L-[14C(U)]arginine at 37°C for 20 min before assay. C: NOS activity in membrane (Memb) and soluble fractions [supernatant (Sup)] of rabbit skeletal muscle. Aliquots of membrane or soluble proteins were incubated with 3.1 mM L-[14C(U)]arginine for 20 min before assay in absence or presence of 2 mM EDTA. D: inhibition of NOS activity by arginine analogs [Nomega -monomethyl-L-arginine (L-NMMA) and Nomega -monomethyl-D-arginine (D-NMMA)] and by calmodulin inhibitors (RS-20 and W-7) in presence of 3.1 mM L-[14C(U)]arginine. Effects of calmodulin inhibitors are shown in absence of external calmodulin. E: NOS activity at various L-arginine concentrations. Vmax, maximal velocity of enzyme activity; Km, dissociation constant. Inset, double reciprocal plot of same data.
[View Larger Version of this Image (36K GIF file)]

The NOS activity was completely inhibited by EDTA (Fig. 2C), by arginine analogs, and by calmodulin inhibitors (Fig. 2D). L-NMMA was about 10 times more effective (50% inhibitory concentration = 1.17 µM) than Nomega -monomethyl-D-arginine (50% inhibitory concentration = 11.3 µM). The NOS activity increased with an increase in the L-arginine concentration in a hyperbolic manner, and double reciprocal plot of the rate vs. substrate concentration resulted in a straight line (Fig. 2E). Irrespective of the method of data analysis (linear or nonlinear curve fitting), essentially the same Vmax and dissociation constant (Km) values were obtained for the same set of data. However, the calculated Vmax varied between 18 and 20 pmol · mg-1 · min-1, and the Km for L-arginine varied between 4 and 5 mM in different experiments (n = 4) when membranes from rabbit back and leg muscles were used.

NOS activity in membranes isolated from control and stimulated muscles was measured in the presence of 20 mM L-[14C(U)]-arginine, a substrate concentration that is four to five times higher than the Km for L-arginine. The NOS activity in control, unstimulated TA and EDL muscles was 21.2 ± 3.8 (n = 3) and 18.7 ± 3.5 pmol · mg-1 · min-1 (n = 3), respectively. In contrast, the NOS activity approximately doubled in the contralateral, in vivo stimulated TA and EDL muscles (Fig. 3).

Fig. 3. Change in skeletal muscle NOS activity during electrical stimulation. NOS activity of control TA (cTA) and EDL (cEDL) as well as of control Sol (cSol) muscle membranes were determined in presence of 20 mM L-[14C(U)]arginine as substrate. Data are means ± SE; n = 3 animals. sTA and sEDL, stimulated TA and EDL, respectively.
[View Larger Version of this Image (13K GIF file)]

Immunoblot analysis revealed an increase in skeletal muscle nNOS expression by chronic low-frequency electrical stimulation of TA and EDL muscles (Fig. 4). An antibody directed against the nNOS recognized a 160-kDa protein band in isolated skeletal muscle membranes. Probing the same blots with an anti-eNOS antibody did not reveal the presence of eNOS in the same skeletal muscle samples. This negative result could be due to the low eNOS levels relative to nNOS in these preparations (16).

Fig. 4. Identification of neural-type NOS (nNOS) in skeletal muscle membranes by Western blot. C and S EDL and TA muscle membranes from 1 rabbit were reacted with a polyclonal antibody against human brain NOS. A positive control sample (provided by Transduction Laboratories) was loaded in nNos lane. Prestained molecular weight (MW) markers, myosin, and beta -galactosidase (beta -gal), are shown in lane 6. Similar data were obtained from 2 other rabbits (not shown).
[View Larger Version of this Image (25K GIF file)]

DISCUSSION

Because nNOS is expressed in type II, fast-twitch skeletal muscle fibers but not in type I, slow-twitch fibers (15), one would expect that a fast-to-slow fiber transformation will result in a decreased expression of nNOS. However, under our experimental conditions an increased level of nNOS expression occurred. This was shown by both activity measurements and immunoblot analysis. The chronic low-frequency electrical stimulation employed in this study represents a well-established method for studying the effects of increased function on expression of specific genes in muscle (21). Many different structural and metabolic changes occur under these conditions (17, 21). A consistent finding has been the fast-to-slow muscle fiber transformation (7, 14, 17, 24, 26, 27). In our experiments this was indicated by increased expression of type I and IIA MHC isoforms, by decreased expression of the type IID MHC isoform, and by the reduction of Vmax in single fibers. Taken together, these data suggest that nNOS expression in skeletal muscle is regulated by changes in muscle activity. However, this regulation does not necessarily follow the fast-twitch and slow-twitch pattern during the dynamic phase of phenotype transformation. There is no evidence to show that fiber transformation and increased NOS activity are functionally related.

Recent data suggest that expression of nNOS in the central nervous system and other tissues such as skeletal muscle is not invariable. In the central nervous system a transient induction of nNOS occurs during synaptogenesis (20). The expression of nNOS is also induced during pregnancy and 17beta -estradiol treatment in the uterus, brain, and the skeletal muscles (30). Our data, by showing that an increase in muscle activity regulates the expression of the nNOS gene in skeletal muscle, further support the notion that nNOS is an inducible enzyme. It is suggested that the nNOS expression in skeletal muscle is under hormonal and metabolic control and the activity of this enzyme in muscle, just like in the central nervous system, may be different under different physiological and pathological conditions.

NO generated in skeletal muscle may have important physiological functions. For example, by inhibition of the ryanodine receptor calcium-release channel (18), NO may decrease calcium release from sarcoplasmic reticulum, depress the contractile force, and decrease the efficiency of excitation contraction coupling (15, 22). NO also increases muscle metabolism by regulating glucose uptake (3) and, as we have shown here, an increase in contractile and metabolic activities during forced stimulation induces the nNOS that produces NO. In addition to these autocrine effects, a portion of the NO produced by muscle fibers may diffuse to neighboring vascular smooth muscle cells and affect them in a paracrine fashion. This is feasible considering that NO is released from contracting muscle and that this release is increased by stimulation of the muscle (3, 15).

On the other hand, excess NO production can have deleterious effects on muscle. It may further decrease the efficiency of excitation-contraction coupling in fast-twitch fibers, in which this coupling is already less efficient than in slow muscles (15, 22). Just like in the nervous system, increased production of NO and reactive oxygen radicals may favor peroxynitrite formation and cell destruction (6, 8). Recent data showing that NO can induce apoptosis in skeletal muscle myoblasts (25) further support the notion that excess NO in the absence of efficient trapping mechanisms may be deleterious to skeletal muscle. However, evidence indicating that NO produced by a constitutive NOS has harmful effects in muscle has not been presented.

ACKNOWLEDGEMENTS

We are grateful to Dr. J. David Johnson for the calmodulin inhibitors and many helpful suggestions and comments. The technical assistance of Sherri Mazetis, Wenrong Wu, and Dr. Laszlo P. Vaghy is appreciated.

FOOTNOTES

   This work was supported by a grant from the Muscular Dystrophy Association (P. L. Vaghy), a Seed Grant from the Ohio State University (P. L. Vaghy), and by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant AR-39652 (P. J. Reiser).

Address for reprint requests: P. L. Vaghy, Dept. of Medical Biochemistry, The Ohio State Univ., 466 Hamilton Hall, 1645 Neil Ave., Columbus, OH 43210-1218 (E-mail: pvaghy{at}magnus.acs.ohio-state.edu).

Received 15 March 1996; accepted in final form 20 November 1996.

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