- Email: [email protected]
Muscle cells from mdx mice have an increased susceptibility to oxidative stress
Neuromuscular Disorders 8 (1998) 14–21
Muscle cells from mdx mice have an increased susceptibility to oxidative stress Thomas A. Rando a , b ,*, Marie-Helene Disatnik a , b, Yip Yu a , b, Alexa Franco a , b
b
a Department of Veterans Affairs, Palo Alto, CA, USA Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, CA 94305, USA
Received 20 August 1997; revised version received 3 October 1997; accepted 8 October 1997
Abstract Several lines of evidence suggest that free radical mediated injury and oxidative stress may lead to muscle necrosis in the muscular dystrophies, including those related to defects in the dystrophin gene. We have examined muscle cell death using an in vitro assay in which the processes that lead to myofiber necrosis in vivo may be amenable to investigation in a simplified cell culture system. Using myotube cultures from normal and dystrophin-deficient (mdx) mice, we have examined the susceptibilities of the cells to different metabolic stresses. Dystrophin-deficient cells were more susceptible to free radical induced injury when compared to normal cells, but the two populations were equally susceptible to other forms of metabolic stress. The differential response appeared to be specifically related to dystrophin expression since undifferentiated myoblasts (which do not express dystrophin) from normal and mdx mice were equally sensitive to oxidative stress. Thus, the absence of dystrophin appears to render muscle specifically more susceptible to free radical induced injury. These results support the hypothesis that oxidative stress may lead to myofiber necrosis in these disorders. Elucidating the mechanisms leading to cell death may help to explain the variabilities in disease expression that are seen as a function of age, among different muscles, and across species in animals with muscular dystrophy due to dystrophin deficiency. 1998 Elsevier Science B.V. Keywords: Muscular dystrophy; mdx; Dystrophin; Oxidative stress; Cell death; Myotubes
1. Introduction The most well characterized dystrophies are due to abnormalities of the dystrophin gene. Defects in the dystrophin gene that result in dystrophin deficiency cause muscular dystrophies in humans (Duchenne muscular dystrophy), mice (the mdx strain), and other species [1,2]. The absence of dystrophin is associated with necrotic muscle cell death, and yet the mechanism by which dystrophin protects the cell from this process is unknown [3]. One of the hallmarks of the muscular dystrophies due to dystrophin deficiency is that the degree to which different muscles are affected varies. In each species, certain muscles are severely affected and other muscles are spared the
* Corresponding author. Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, CA 943055235, USA.
0960-8966/98/$19.00 1998 Elsevier Science B.V. All rights reserved PII S0960-8966 (97 )0 0124-7
degenerative process [1,2,4,5]. Many factors have been proposed to explain these differences such as fiber size, fiber type composition, and embryological origin [6–8]. In addition, the extent of active necrosis can vary tremendously with age, a feature that has been well characterized in different animals and particularly in the mdx mouse [2,9]. Thus, dystrophin deficiency is not a disorder of uniform, inevitable muscle necrosis; rather, it is a disorder of susceptibility to necrosis, and that susceptibility may vary over space and time. This issue is further highlighted by the fact that normal muscle may undergo necrotic changes characteristic of muscular dystrophies when exposed to severe oxidative stress [10–13]. In this regard, dystrophin-deficient and normal muscle may fall on a spectrum of susceptibility to specific forms of injury, with normal muscle being relatively more resistant. Understanding that susceptibility (or resistance) and how it differs between normal and dystrophin-deficient muscles, or how it varies among different dystrophin-deficient muscles, is a key to elucidating the
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
primary pathogenetic mechanisms leading to muscle cell necrosis in these disorders. Because of similarities of the pathology of dystrophin deficiency to that of muscle injury due to oxidative stress [10–13], it has been postulated that the cell death in muscular dystrophies may be due to free radical mediated injury [14,15]. Indeed, there is evidence from in vivo studies that free radical injury to muscle cell membrane may underlie the primary degenerative process of muscle cell death [14,16–19]. According to this free radical hypothesis, the necrosis of muscle in an animal with a muscular dystrophy would occur because their muscles would be more susceptible to oxidative injury than would normal muscle. It would be predicted that, at a given level of oxidative stress, dystrophin-deficient muscle would undergo necrotic degeneration whereas normal muscle, exposed to the same level of stress, would remain intact. In this report, we present data showing that dystrophin deficiency is associated with an increased susceptibility of muscle to oxidative stress. These findings are consistent with predictions of the free radical hypothesis, and the in vitro assay system presented here should allow tests of this and other theories of the cause of muscle necrosis in muscular dystrophies.
2. Materials and methods 2.1. Animals Mice (strains C57 (C57BL/10SnJ) and mdx (C57BL/ 10ScSn-mdx) were obtained from the Jackson Laboratory (Bar Harbor, ME). Animals were handled in accordance with guidelines of the Administrative Panel on Laboratory Animal Care of Stanford University. 2.2. Muscle cell cultures Primary cultures of skeletal muscle were prepared, and enriched myoblast cultures were derived as described previously [20]. Using these conditions, cultures were purified to greater than 99% myogenic cells, and therefore there was no contribution of non-myogenic cells in these studies [20]. To assess growth rates, 5 × 104 cells were plated in 35 mm laminin-coated dishes in growth medium (GM) consisting of Ham’s F-10 nutrient mixture (GIBCO BRL, Gaithersburg, MD) supplemented with 20% fetal bovine serum (HyClone Laboratories, Inc., Logan, UT), 2.5 ng/ml bFGF (Promega Corp., Madison, WI), penicillin and streptomycin. The number of cells in each dish was determined at different intervals by hemacytometer counts after the cells were suspended by trypsinization. All conditions were tested in triplicate cultures. The population doubling times of the cells on different substrates were determined during periods when cells were in log-phase growth. Myoblast cultures were induced to differentiate into myotubes by maintaining the cells in mitogen-poor differentia-
15
tion medium (DM) consisting of Dulbecco’s modified Eagle’s medium (GIBCO) supplemented with 2% horse serum (HyClone), penicillin and streptomycin [20]. To assess morphologic differentiation, 5 × 104 cells were plated in 35 mm laminin-coated dishes in growth medium. After 24 h, the growth medium was replaced by DM. The cells were maintained in DM with daily medium changes, and all measurements were obtained from triplicate cultures. The fusion index was determined as the ratio of the number of nuclei in myotubes (cells with ≥3 nuclei) to the total number of nuclei. Ten randomly chosen fields from each dish were counted at a magnification of 250 × . 2.3. Immunoblotting Cell extracts were prepared in a buffer consisting of 50 mM Tris–HCl, 5 mM EDTA, 5 mM EGTA, 20 mg/ml leupeptin, 20 mg/ml aprotinin, 100 mg/ml PMSF, and 50 mM DTT. For each sample, the protein content was determined using the Bio-Rad protein assay. Samples (20 mg protein/ lane) of cell extracts were run on 5% SDS-polyacrylamide gels, transferred to nitrocellulose membranes (0.45 mm; Schleicher and Schuell, Keene, NH), and probed with a mouse monoclonal antibody to dystrophin (Mandys 8; Sigma Chemical, St. Louis, MO) at a 1:400 dilution. The specific antibody reaction was detected by an enhanced chemiluminescence system (Amersham Corp., Arlington Heights, IL). 2.4. Immunocytochemistry Myotube cultures were fixed in 2% formaldehyde in PBS for 5 min at room temperature followed by 100% methanol at −20°C for 5 min. The cells were then rinsed in PBS. All further incubations were carried out at room temperature and all rinses and dilutions were with a blocking solution consisting of 5% horse serum and 0.5% Triton-X 100 in PBS. An initial blocking step was performed with this solution for 30 min. A mouse monoclonal antibody to dystrophin (Dys1 - Novocastra Laboratories; Vector Laboratories, Burlingame, CA) was applied neat for 30 min. After rinsing, a Texas Red-coupled secondary antibody (Cappel Research Products, Durham, NC) was applied for 30 min at a dilution of 1:250. The cells were examined and photographed with a Zeiss Axioskop microscope. 2.5. Cell survival assay Myoblasts were plated in GM into 24 well plates at a density of 7.5 × 104 cells/well. After 1 day, the medium was changed to DM. Cells were differentiated for 4–5 days (myotube formation began after day 2), and cellular toxins were added to each well for 12–18 h as indicated. Cell survival was determined using a kit for measuring lactate dehydrogenase (LDH) activity according to the instructions of the manufacturer (Promega, Madison, WI). Briefly,
16
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
cellular LDH was used as a measure of the number of surviving cells in each well. After the incubation period, the supernatant was removed, the remaining cells were rinsed, and the cellular LDH activity was measured. Cell death was calculated as a percentage, with 0% cell death being the value of cellular LDH in wells with no added toxin, and 100% cell death being the value of cellular LDH remaining after addition of 0.9% SDS to the culture medium just prior to the end of the incubation period. Thus, for each cell population, there were internal positive and negative controls for the assay itself. All data are from assays done in triplicate. During initial studies to validate the use of the LDH assay, cell death was corroborated in parallel myoblast cultures by hemacytometer counts of total and viable cell numbers using trypan blue exclusion after trypsinization, and there was an excellent agreement between the two methods. It was also routine to estimate cell death by visual inspection of the cultures at the end of the incubation period as a further corroboration (estimating to the closest 10% over the range of 0% (untreated cultures) to 100% cell death), and these estimates paralleled the LDH results as well. For myotube cultures, hemacytometer counts are not accurate, so corroboration of LDH assay results was always done by visual inspection of the cultures before the assay. There was never a discordance between the general level of cell death by the two methods, but of course estimates of cell death by visual inspection were less accurate than those by the LDH assay. 2.6. Cellular toxins Cellular toxins were divided into ‘pro-oxidants’ (those whose mechanism of toxicity was primarily by promoting oxidative injury) and ‘non-oxidants’ (those whose mechanism of injury was not primarily due to free radical injury). In each case, dose-response curves were done and the levels of cell death were determined over time. The incubation time chosen for these studies was in the range of 12–18 h because this resulted in a ‘steady-state’ level of cell death in response to the one-time addition of each toxin to the medium across the concentration ranges listed below. All reagents were purchased from Sigma Chemical (St. Louis, MO) unless otherwise indicated. Pro-oxidants tested were: H2O2: 0.01–31.6 mM; H2O2 generates hydroxyl radicals by metal-catalyzed reactions (Fenton chemistry) [21]. Paraquat: 0.01–31.6 mM; paraquat generates superoxide intracellularly [22]. Menadione: 0.1 mM–1 mM; menadione is a quinone that generates free radicals intracellularly by undergoing repeated redox cycles [23]. AAPH (2,2′-amino azo bis amidopropane hydrochloride; Wako Chemicals, Richmond, VA): 0.1–100 mM;
AAPH generates peroxyl radicals that can initiate lipid peroxidation [24]. SNP (sodium nitroprusside): 0.01–31.6 mM; SNP generates nitric oxide [25] which combines with endogenous superoxide to form the potent reactive oxygen species, peroxynitrite [26]. Non-oxidants tested were: Staurosporine: 1 nM–1 mM; staurosporine has been shown to promote apoptotic cell death in a variety of cell types by the inhibition of protein kinases [27]. A23187: 0.1–100 mg/ml; the primary mechanism of toxicity is via rapid rises in intracellular calcium and activation of proteases, phospholipases, and endonucleases [28]. CCCP (carbonyl cyanide m-chlorophenylhydrazone): 0.1–100 mM; CCCP is an inhibitor of mitochondrial respiration. The primary mechanism of toxicity of mitochondrial inhibitors is via reduction of cellular ATP levels, and the resulting cellular injury is independent of oxidant injury [29]. 2.7. Statistical analysis Comparisons between samples were done by analysis of variance (ANOVA). Differences were considered statistically significant at the P , 0.05 level.
3. Results Undifferentiated skeletal myoblasts were derived from neonatal mice [20] of both the C57 and the mdx strains. Myoblasts from C57 and mdx muscle behaved similarly in assays of cell growth (population doubling times were 19.4 ± 0.8 h for C57 myoblasts, and 18.9 ± 0.5 h for mdx myoblasts) and differentiation (the fusion indices were 0.38 ± 0.04 in C57 cultures and 0.39 ± 0.02 in mdx cultures after 3 days in DM). When grown in mitogen-poor medium, skeletal myoblasts differentiated into multinucleated myotubes (Fig. 1) which expressed many of the morphological and biochemical properties of mature muscle cells in vivo (e.g., expression of myosin heavy chain, skeletal muscle actin, creatine phosphokinase), except for the fact that mdx myotubes did not express dystrophin (Fig. 1). We exposed both C57 and mdx myotube cultures to prooxidant compounds to compare their susceptibilities to oxidative stress. Fig. 2 shows dose-response curves of cell death as a function of concentration of two compounds that induce oxidative stress, the reactive oxygen species H2O2 and the superoxide-generating compound, paraquat. For both compounds, mdx myotubes were more susceptible to the oxidative injury than were normal myotubes as evidenced by greater cell death across a wide concentration range. In order to compare the susceptibility of the two
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
17
Fig. 1. Differentiation of C57 and mdx muscle cells in culture. (A) Myotube formation by C57 and mdx muscle cells. Myoblast cultures derived from C57 and mdx mice were induced to differentiate by culturing in low serum medium. After 5 days, myoblast elongation and fusion lead to multinucleated myotube formation in each culture. The top panels show phase contrast images of the myotube cultures. The bottom panels show immunofluorescent staining for dystrophin in the two populations. Dystrophin expression was detected only in the C57 myotubes. (B) Time course of expression of dystrophin in myotube cultures. Cultures of muscle cells from both C57 and mdx mice were maintained in DM, and each day cultures were analyzed by immunoblot analysis for the expression of dystrophin. Dystrophin expression increased during the first several days in C57 cultures, and plateaued after 3–4 days. No dystrophin expression was observed in the mdx cultures.
Fig. 2. Dystrophin-deficient myotubes are more susceptible to oxidative injury than are normal myotubes. Both C57 and mdx myotube cultures were subjected to oxidative stress by the addition of the pro-oxidant compounds (A) H2O2 or (B) paraquat to the culture medium. Sixteen hours after exposure to different concentrations of the pro-oxidants, the extent of cell death was determined by the LDH assay. In both cases, mdx myotubes exhibited greater sensitivity to oxidative stress than did C57 myotubes. Data are expressed as mean ± SD (n = 4).
18
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
Fig. 3. Comparisons of susceptibilities of C57 and mdx muscle cells to oxidative stress and non-oxidative metabolic stresses. Experiments were done as in Fig. 2, and the extent of cell death was compared at concentrations of the toxins that resulted in sub-maximal cell death. Data are expressed as mean ± SD (n = 4). (A) Susceptibility of C57 and mdx myotubes to oxidative stress. The pro-oxidants (see Section 2) were tested at the following concentrations: 0.316 mM H2O2; 5 mM menadione; 2 mM paraquat; 16.5 mM AAPH; 1 mM SNP. Despite differences in mechanisms of producing oxidative stress among the different toxins, mdx myotubes were more susceptible than C57 myotubes to all of the compounds tested. (B) Susceptibility of C57 and mdx myotubes to non-oxidant toxins (see Section 2). The non-oxidants were tested at the following concentrations: 0.01 mM staurosporine; 2 mg/ml A23187; 10 mM CCCP. The two populations of myotubes were equally susceptible to all of the non-oxidant compounds tested.
cell populations, we chose concentrations of the toxins that produced sub-maximal levels of cell death in mdx cultures and detectable levels of cell death in C57 cultures. Fig. 3A summarizes the results of these and similar experiments with a variety of pro-oxidant compounds, each of which induces oxidative stress by a different mechanism (see Section 2). In each case, the susceptibility of dystrophin-deficient myotubes to oxidative injury was significantly greater than that of the normal myotubes. Although these results demonstrated that dystrophin-deficient muscle was more sensitive than normal muscle to oxidative stress, we subjected the cells to non-oxidant forms of injury to test the possibility that dystrophin deficient muscle was just more susceptible to any form of metabolic stress. Thus, C57 and mdx myotube cultures were exposed to toxins whose mechanisms of injury are not via free radical mechanisms (see Section 2). C57 and mdx myotube cultures were equally sensitive to the toxic effects of
these agents at each concentration tested, and the effects of the dose of each toxin that produced approximately 50% cell death is shown (Fig. 3B). Certainly, any significant difference in susceptibility of the cells to these toxins would be apparent at these intermediate concentrations. These results suggest that dystrophin-deficient muscle cells are specifically more susceptible to oxidative stress, and not simply more susceptible to any form of metabolic stress that might cause cellular injury. C57 and mdx mice are from the same genetic background, and differ genetically by the presence of a mutation in the dystrophin gene in the mdx strain [30]. To confirm that the difference in susceptibility to oxidative stress of the two strains was indeed due to the difference in dystrophin expression, we compared the susceptibility of undifferentiated myoblasts from the two strains to oxidative stress. Since dystrophin is not expressed in myoblasts, the two populations should be identical biochemically, and we predicted that they would be equally susceptible to any form of injury. Indeed, Fig. 4 shows that the C57 and mdx myoblasts were equally susceptible to the toxic effects of paraquat. There was no difference in susceptibility to every metabolic toxin tested, including the other pro-oxidants H2O2 and menadione, as well as the control toxins staurosporine and A23187. Finally, as a further indication that dystrophin expression confers a certain level of resistance to oxidative injury, we compared the susceptibilities of myotubes from four normal strains of mice: C57, C3H, SJL, and BALB/c. All four strains express dystrophin and have no intrinsic muscle disorder. Among the four strains tested, there were no differences in their susceptibilities to pro-oxidants (Fig. 5), nor was there any difference in susceptibilities to non-oxidants. This was true over a range of concentrations that resulted in anywhere from 25% to 75% cell death, the most sensitive
Fig. 4. Susceptibilities of C57 and mdx myoblasts to oxidative stress. Undifferentiated myoblasts were maintained in growth medium and exposed to different concentrations of paraquat for 18 h. Cell death was measured using the LDH assay. Data are presented as mean ± SD (n = 5). Unlike myotube cultures, mdx myoblasts were not more susceptible to oxidative stress than were C57 myoblasts.
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
Fig. 5. Susceptibilities of myotubes from different normal strains of mice to oxidative stress. In addition to the C57 strain, which is the parental strain of the mdx, we tested the relative susceptibilities of myotube cultures from different dystrophin-positive strains to oxidative stress. These data show the results of the addition of either (A) menadione (10 mM) or (B) paraquat (5 mM) to the culture medium of myotube cultures from the C57, C3H, BALB/c, and SJL strains of mice. Cell death was measured after 12 h of exposure, and data are presented as mean ± SD (n = 4). There was no significant difference between the results of any of the strains; all were equally susceptible to oxidative stress.
range for detecting any differences between the cell populations. Thus, among five independently derived primary muscle cell cultures, only the dystrophin-deficient myotubes showed an increased susceptibility to oxidative stress.
4. Discussion Selective muscle involvement is a hallmark of the muscular dystrophies, indicating that genetic defects present in all muscles do not affect all muscles equally [1,2,5]. In muscular dystrophies due to dystrophin deficiency, there are examples of specific muscle sparing in humans and
19
other animal species. In particular, in the mdx mouse there is no evidence of muscle necrosis for the first several weeks postnatally [9]. There follows a period of widespread necrosis over the next 6–8 weeks, after which there is minimal necrosis in limb muscles for the remainder of the adult life. By contrast, there is also evidence in all species examined that certain muscles are severely affected [1,2,5]. Clearly, epigenetic mechanisms play an important role in determining whether a muscle will succumb to the degenerative process, when the degeneration will begin (or end), and how severely a muscle will be affected. The relative susceptibility or resistance of each muscle will account for phenotypic variability. Examining cellular susceptibility in vivo is particularly complex because of the many uncontrolled variables that may affect cell survival or cell death. For this reason, cellular susceptibility is commonly studied in culture where pharmacologic agents are used to mimic physiologic stresses, such as oxidative stress, that may occur in vivo [31–33]. We have taken this approach by using a purified population of myogenic cells. While lacking the complexities of mature muscle tissue, these studies allow analysis using a single end-point that occurs both in vivo and in vitro, namely cell death. Our results suggest that dystrophin deficiency renders muscle susceptible to oxidative stress, but not merely to any form of metabolic stress. The fact that both extreme exercise [12,13] and muscle ischemia [10,11] can generate lesions that resemble dystrophic changes, and that such lesions are mediated by oxidative injury [34,35], supports the hypothesis that a muscular dystrophy could be a consequence of an increased susceptibility to this form of metabolic stress. Furthermore, similarities between the pathology of muscular dystrophies and the myopathy of vitamin E deficiency, a condition of increased susceptibility to oxidative stress, have long been noted [14]. Recently, we have also documented a muscular dystrophy caused by a genetic alteration in free radical metabolism. In a transgenic mouse that overexpresses the enzyme Cu,Zn-superoxide dismutase [36], we have found a profound muscular dystrophy that begins between 2 and 4 months of age (unpublished observations). Overexpression of this enzyme has been reported to sensitize cells to oxidative stress by producing elevated levels of H2O2 [33,37]. As a model for a muscular dystrophy specifically due to oxidative stress, this mouse strain should prove to be very informative. The role of free radical mediated injury in the pathogenesis of the muscular dystrophies has been extensively studied in vivo [14]. Working with mammalian and avian dystrophic muscle, investigators have demonstrated increased lipid peroxidation [14,38], increased oxidation of proteins [16,17], and induction of antioxidant enzymes [39,40]. We have studied the role of oxidative injury in the mdx mouse in vivo by focusing on the pre-necrotic state, since the onset of muscle degeneration does not begin until about 3 weeks of age. We have found an increase in lipid
20
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
peroxidation in mdx muscle preceding the onset of muscle necrosis [19]. The free radical hypothesis complements many current ideas on the pathogenesis of the disease [3]. The ‘mechanical damage hypothesis’, the ‘leaky membrane hypothesis’, and the ‘calcium hypothesis’ all address some of the cardinal features of dystrophic muscle, there is evidence to support aspects of each of these theories [41–43], and they are not mutually exclusive. Certainly, there appears ultimately to be a disruption of membrane integrity with subsequent calcium influx leading to cell necrosis, but the biochemical process that initiates this chain of events is unspecified and the resistance of certain muscles to the degenerative process remains unexplained. The free radical hypothesis suggests just such a biochemical cascade: normal cellular metabolism produces free radicals, free radicals cause lipid peroxidation, lipid peroxidation causes membrane breakdown and cell necrosis. According to the hypothesis and as suggested by our data, the absence of dystrophin would be associated with an increased susceptibility to oxidative injury, whereas the membrane of normal muscle would be more resistant. Cellular antioxidant capacity differs among muscles and as a function of age [44,45], and such changes in cell protective mechanisms may explain how a disease resulting from a constant genetic defect could be manifested differently over space and time [18]. As dystrophin is not known to have any antioxidant properties, the question arises as to how dystrophin may function to protect the cell against free radical injury, and, conversely, how the absence of dystrophy could render a cell susceptible to free radical injury. The recent demonstration of the association of the dystrophin complex with an enzyme, namely nitric oxide synthase (NOS), involved in free radical metabolism is intriguing [46,47]. This is the first demonstration of the association of dystrophin with a cellular metabolic pathway, and that pathway happens to involve free radical metabolism, namely the generation of the free radical nitric oxide (NO). How this association, and its disruption in dystrophin-deficient muscle [46,47], may relate to the pathogenesis of muscular dystrophies remains to be determined. Alterations in NOS activity could alter NO levels in the cell, and NO has been shown to have both protective and destructive actions in cellular oxidative injury [48]. We are currently exploring the role of NO in muscle cell oxidative metabolism and free radical injury. The possibility that oxidative stress may play an important role in the pathogenesis of muscle degeneration in the dystrophies could have profound therapeutic consequences. Clinical trials of antioxidants have been discouraging in Duchenne muscular dystrophy [49,50], but these have always involved patients long after the onset and progression of the degeneration. Our results suggest that oxidative injury might contribute to the very first pathologic change, namely cell death, and that antioxidant therapy could be effective only if started early enough to prevent muscle cell necrosis. The mdx mouse is an ideal model system for
such trials, since the onset of muscle necrosis occurs after approximately 3 weeks of age [3]. The benefit of early intervention with antioxidant therapy could be tested in the mdx mouse to determine if such trials would be warranted in humans with Duchenne muscular dystrophy.
Acknowledgements The authors would like to thank Dr. Grace Pavlath of Emory University for supplying myoblast cultures from BALB/c and SJL strains. This work was supported in part by a Howard Hughes Summer Fellowship from the Department of Biological Sciences at Stanford University to AAF and by grants from the Department of Veterans Affairs, the Muscular Dystrophy Association, and the American Academy of Neurology Education and Research Foundation to TAR. References [1] Emery AEH. Duchenne muscular dystrophy. New York: Oxford University Press, 1993. [2] Hoffman EP, Gorospe JRM. The animal models of Duchenne muscular dystrophy: windows on the pathophysiological consequences of dystrophin deficiency. Curr Top Memb 1991;38:113–154. [3] McArdle A, Edwards RHT, Jackson MJ. How does dystrophin deficiency lead to muscle degeneration? – Evidence from the mdx mouse. Neuromusc Disord 1995;5:445–456. [4] Valentine BA, Cooper BJ. Canine X-linked muscular dystrophy: selective involvement of muscles in neonatal dogs. Neuromusc Disord 1991;1:31–38. [5] Stedman HH, Sweeney HL, Shrager JB. et al. The mdx mouse diaphragm reproduces the degenerative changes of Duchenne muscular dystrophy. Nature 1991;352:536–538. [6] Boland B, Himpens B, Denef JF, Gillis JM. Site-dependent pathological differences in smooth muscles and skeletal muscles of the adult mdx mouse. Muscle Nerve 1995;18:649–657. [7] Minetti C, Ricci E, Bonilla E. Progressive depletion of fast alphaactinin-positive muscle fibers in Duchenne muscular dystrophy. Neurology 1991;41:1977–1981. [8] Karpati G, Carpenter S, Prescott S. Small-caliber skeletal muscle fibers do not suffer necrosis in mdx mouse dystrophy. Muscle Nerve 1988;11:795–803. [9] Tanabe Y, Esaki K, Nomura T. Skeletal muscle pathology in X chromosome-linked muscular dystrophy (mdx) mouse. Acta Neuropathol 1986;69:91–95. [10] Mendell JR, Engel WK, Derrer EC. Duchenne muscular dystrophy: functional ischemia reproduces its characteristic lesions. Science 1971;172:1143–1145. [11] Engel WK. Duchenne muscular dystrophy: a histologically based ischemia hypothesis and comparison with experimental ischemia myopathy. In: Pearson CM, Mostofi FK, editors. The striated muscle. Baltimore: Williams and Wilkins, 1973:453–472. [12] Vihko V, Rantamaki J, Salminen A. Exhaustive physical exercise and acid hydrolase activity in mouse skeletal muscle. Histochemistry 1978;57:237–249. [13] Irintchev A, Wernig A. Muscle damage and repair in voluntarily running mice: strain and muscle differences. Cell Tissue Res 1987;249:509–521. [14] Murphy ME, Kehrer JP. Oxidative stress and muscular dystrophy. Chem-Biol Interact 1989;69:101–178.
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21 [15] Brown RH. Free radicals, programmed cell death, and muscular dystrophy. Curr Opin Neurol 1995;8:373–378. [16] Hauser E, Hoger H, Bittner R, Widhalm K, Herkner K, Lubec G. Oxyradical damage and mitochondrial enzyme activities in the mdx mouse. Neuropediatrics 1995;26:260–262. [17] Haycock JW, MacNeil S, Jones P, Harris JB, Mantle D. Oxidative damage to muscle protein in Duchenne muscular dystrophy. NeuroReport 1996;8:357–361. [18] Ragusa FJ, Chow CK, St Clair DK, Porter JD. Extraocular, limb, and diaphragm muscle group-specific antioxidant enzyme activity patterns in control and mdx mice. J Neurol Sci 1996;139:180–186. [19] Rando TA, Disatnik M-H, Dhawan J, Yu Y, Whirl MM. Oxidative injury precedes muscle cell necrosis in dystrophin-deficient (mdx) mice. Neurology 1997;48:A442. [20] Rando TA, Blau HM. Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol 1994;125:1275–1287. [21] Lloyd RV, Hanna PM, Mason RP. The origin of the hydroxyl radical oxygen in the Fenton reaction. Free Rad Biol Med 1997;22:885–888. [22] Krall J, Bagley AC, Mullenbach GT, Hallewell RA, Lynch RE. Superoxide mediates the toxicity of paraquat for cultured mammalian cells. J Biol Chem 1988;263:1910–1914. [23] Thor H, Smith MT, Hatzell P, Bellomo G, Jewell SA, Orrenius S. The metabolism of menadione (2-methyl-1,4-naphthoquinone) by isolated hepatocytes. A study of the implications of oxidative stress in intact cells. J Biol Chem 1982;257:12419–12425. [24] Dooley MM, Sano N, Kawashima H, Nakamura T. Effects of 2,2′azobis (2-amidinopropane) hydrochloride in vivo and protection by vitamin E. Free Rad Biol Med 1990;9:199–204. [25] Feelisch M, Noack EA. Correlation between nitric oxide formation during degradation of organic nitrates and activation of guanylate cyclase. Eur J Pharmacol 1987;139:19–30. [26] Beckman JS, Beckman TW, Chen J, Marshall PA, Freeman BA. Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide. Proc Natl Acad Sci USA 1990;87:1620–1624. [27] Jacobson MD, Raff MC. Programmed cell death and Bcl-2 protection in very low oxygen. Nature 1995;374:814–816. [28] Nicotera P, Bellomo G, Orrenius S. Calcium-mediated mechanisms in chemically induced cell death. Annu Rev Pharmacol Toxicol 1992;32:449–470. [29] Weinberg JM, Davis JA, Abarzua M, Kaini T, Kunkel R. Protection by glycine of proximal tubules from injury due to inhibitors of mitochondrial ATP production. Am J Physiol 1990;258:C1127–C1140. [30] Sicinski P, Geng Y, Ryder-Cook AS, Barnard EA, Darlison MG, Barnard PJ. The molecular basis of muscular dystrophy in the mdx mouse: a point mutation. Science 1989;244:1578–1580. [31] Zhong L-T, Sarafian T, Kane DJ. et al. bcl-2 Inhibits death of central neural cells induced by multiple agents. Proc Natl Acad Sci USA 1993;90:4533–4537. [32] Bonfoco E, Krainc D, Ankarcrona M, Nicotera P, Lipton SA. Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-d-aspartate or nitric oxide/superoxide in cortical cell cultures. Proc Natl Acad Sci USA 1995;92:7162–7166.
21
[33] Amstad P, Moret R, Cerutti P. Glutathione peroxidase compensates for the hypersensitivity of Cu,Zn-superoxide dismutase overproducers to oxidant stress. J Biol Chem 1994;269:1606–1609. [34] Radak Z, Asano K, Inoue M. et al. Superoxide dismutase derivative reduces oxidative damage in skeletal muscle of rats during exhaustive exercise. J Appl Physiol 1995;79:129–135. [35] Fantini GA, Yoshioka T. Deferoxamine prevents lipid peroxidation and attenuates reoxygenation injury in postischemic skeletal muscle. Am J Physiol 1993;264:H1953–H1959. [36] Epstein CJ, Avraham KB, Lovett M. et al. Transgenic mice with increased Cu/Zn-superoxide dismutase activity: animal model of dosage effects in Down syndrome. Proc Natl Acad Sci USA 1987;84:8044–8048. [37] Amstad P, Peskin A, Shah G. et al. The balance between Cu,Znsuperoxide dismutase and catalase affects the sensitivity of mouse epidermal cells to oxidative stress. Biochem 1991;30:9305–9313. [38] Mizuno Y. Changes in superoxide dismutase, catalase, glutathione peroxidase, and glutathione reductase activities and thiobarbituric acid-reactive product levels in early stages of development in dystrophic chickens. Exp Neurol 1984;84:58–73. [39] Burr IM, Asayama K, Fenichel G. Superoxide dismutase, glutathione peroxidase, and catalase in neuromuscular disease. Muscle Nerve 1987;10:150–154. [40] Austin L, de Niese M, McGregor A, Arthur H, Gurushinghe A, Gould MK. Potential oxyradical damage and energy status in individual muscle fibres from degenerating muscle diseases. Neuromusc Disord 1992;2:27–33. [41] Petrof BJ, Shrager JB, Stedman HH, Kelly AM, Sweeney HL. Dystrophin protects the sarcolemma from stresses developed during muscle contraction. Proc Natl Acad Sci USA 1993;90:3710–3714. [42] Turner PR, Fong P, Denetclaw WF, Steinhardt RA. Increased calcium influx in dystrophic muscle. J Cell Biol 1991;115:1701–1712. [43] Rowland LP. Biochemistry of muscle membranes in Duchenne muscular dystrophy. Muscle Nerve 1980;3:3–20. [44] Powers SK, Criswell D, Lawler J. et al. Influence of exercise and fiber type on antioxidant enzyme activity in rat skeletal muscle. Am J Physiol 1994;266:R375–R380. [45] Oh-Ishi S, Kizaki T, Yamashita H. et al. Alterations of superoxide dismutase iso-enzyme activity, content, and mRNA expression with aging in rat skeletal muscle. Mech Ageing Dev 1995;84:65–76. [46] Brenman JE, Chao DS, Xia H, Aldape K, Bredt DS. Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell 1995;82:743– 752. [47] Chang W-J, Iannaccone ST, Lau KS. et al. Neuronal nitric oxide synthase and dystrophin-deficient muscular dystrophy. Proc Natl Acad Sci USA 1996;93:9142–9147. [48] Darley-Usmar V, Wiseman H, Halliwell B. Nitric oxide and oxygen radicals: a question of balance. FEBS Lett 1995;369:131–135. [49] Roelofs RI, de Arango GS, Law PK, Kinsman D, Buchanan DC, Park JH. Treatment of Duchenne’s muscular dystrophy with penicillamine. Arch Neurol 1979;36:266–268. [50] Fenichel GM, Brooke MH, Griggs RC. et al. Clinical investigation in Duchenne muscular dystrophy: penicillamine and vitamin E. Muscle Nerve 1988;11:1154–1168.
Muscle cells from mdx mice have an increased susceptibility to oxidative stress Thomas A. Rando a , b ,*, Marie-Helene Disatnik a , b, Yip Yu a , b, Alexa Franco a , b
b
a Department of Veterans Affairs, Palo Alto, CA, USA Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, CA 94305, USA
Received 20 August 1997; revised version received 3 October 1997; accepted 8 October 1997
Abstract Several lines of evidence suggest that free radical mediated injury and oxidative stress may lead to muscle necrosis in the muscular dystrophies, including those related to defects in the dystrophin gene. We have examined muscle cell death using an in vitro assay in which the processes that lead to myofiber necrosis in vivo may be amenable to investigation in a simplified cell culture system. Using myotube cultures from normal and dystrophin-deficient (mdx) mice, we have examined the susceptibilities of the cells to different metabolic stresses. Dystrophin-deficient cells were more susceptible to free radical induced injury when compared to normal cells, but the two populations were equally susceptible to other forms of metabolic stress. The differential response appeared to be specifically related to dystrophin expression since undifferentiated myoblasts (which do not express dystrophin) from normal and mdx mice were equally sensitive to oxidative stress. Thus, the absence of dystrophin appears to render muscle specifically more susceptible to free radical induced injury. These results support the hypothesis that oxidative stress may lead to myofiber necrosis in these disorders. Elucidating the mechanisms leading to cell death may help to explain the variabilities in disease expression that are seen as a function of age, among different muscles, and across species in animals with muscular dystrophy due to dystrophin deficiency. 1998 Elsevier Science B.V. Keywords: Muscular dystrophy; mdx; Dystrophin; Oxidative stress; Cell death; Myotubes
1. Introduction The most well characterized dystrophies are due to abnormalities of the dystrophin gene. Defects in the dystrophin gene that result in dystrophin deficiency cause muscular dystrophies in humans (Duchenne muscular dystrophy), mice (the mdx strain), and other species [1,2]. The absence of dystrophin is associated with necrotic muscle cell death, and yet the mechanism by which dystrophin protects the cell from this process is unknown [3]. One of the hallmarks of the muscular dystrophies due to dystrophin deficiency is that the degree to which different muscles are affected varies. In each species, certain muscles are severely affected and other muscles are spared the
* Corresponding author. Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, CA 943055235, USA.
0960-8966/98/$19.00 1998 Elsevier Science B.V. All rights reserved PII S0960-8966 (97 )0 0124-7
degenerative process [1,2,4,5]. Many factors have been proposed to explain these differences such as fiber size, fiber type composition, and embryological origin [6–8]. In addition, the extent of active necrosis can vary tremendously with age, a feature that has been well characterized in different animals and particularly in the mdx mouse [2,9]. Thus, dystrophin deficiency is not a disorder of uniform, inevitable muscle necrosis; rather, it is a disorder of susceptibility to necrosis, and that susceptibility may vary over space and time. This issue is further highlighted by the fact that normal muscle may undergo necrotic changes characteristic of muscular dystrophies when exposed to severe oxidative stress [10–13]. In this regard, dystrophin-deficient and normal muscle may fall on a spectrum of susceptibility to specific forms of injury, with normal muscle being relatively more resistant. Understanding that susceptibility (or resistance) and how it differs between normal and dystrophin-deficient muscles, or how it varies among different dystrophin-deficient muscles, is a key to elucidating the
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
primary pathogenetic mechanisms leading to muscle cell necrosis in these disorders. Because of similarities of the pathology of dystrophin deficiency to that of muscle injury due to oxidative stress [10–13], it has been postulated that the cell death in muscular dystrophies may be due to free radical mediated injury [14,15]. Indeed, there is evidence from in vivo studies that free radical injury to muscle cell membrane may underlie the primary degenerative process of muscle cell death [14,16–19]. According to this free radical hypothesis, the necrosis of muscle in an animal with a muscular dystrophy would occur because their muscles would be more susceptible to oxidative injury than would normal muscle. It would be predicted that, at a given level of oxidative stress, dystrophin-deficient muscle would undergo necrotic degeneration whereas normal muscle, exposed to the same level of stress, would remain intact. In this report, we present data showing that dystrophin deficiency is associated with an increased susceptibility of muscle to oxidative stress. These findings are consistent with predictions of the free radical hypothesis, and the in vitro assay system presented here should allow tests of this and other theories of the cause of muscle necrosis in muscular dystrophies.
2. Materials and methods 2.1. Animals Mice (strains C57 (C57BL/10SnJ) and mdx (C57BL/ 10ScSn-mdx) were obtained from the Jackson Laboratory (Bar Harbor, ME). Animals were handled in accordance with guidelines of the Administrative Panel on Laboratory Animal Care of Stanford University. 2.2. Muscle cell cultures Primary cultures of skeletal muscle were prepared, and enriched myoblast cultures were derived as described previously [20]. Using these conditions, cultures were purified to greater than 99% myogenic cells, and therefore there was no contribution of non-myogenic cells in these studies [20]. To assess growth rates, 5 × 104 cells were plated in 35 mm laminin-coated dishes in growth medium (GM) consisting of Ham’s F-10 nutrient mixture (GIBCO BRL, Gaithersburg, MD) supplemented with 20% fetal bovine serum (HyClone Laboratories, Inc., Logan, UT), 2.5 ng/ml bFGF (Promega Corp., Madison, WI), penicillin and streptomycin. The number of cells in each dish was determined at different intervals by hemacytometer counts after the cells were suspended by trypsinization. All conditions were tested in triplicate cultures. The population doubling times of the cells on different substrates were determined during periods when cells were in log-phase growth. Myoblast cultures were induced to differentiate into myotubes by maintaining the cells in mitogen-poor differentia-
15
tion medium (DM) consisting of Dulbecco’s modified Eagle’s medium (GIBCO) supplemented with 2% horse serum (HyClone), penicillin and streptomycin [20]. To assess morphologic differentiation, 5 × 104 cells were plated in 35 mm laminin-coated dishes in growth medium. After 24 h, the growth medium was replaced by DM. The cells were maintained in DM with daily medium changes, and all measurements were obtained from triplicate cultures. The fusion index was determined as the ratio of the number of nuclei in myotubes (cells with ≥3 nuclei) to the total number of nuclei. Ten randomly chosen fields from each dish were counted at a magnification of 250 × . 2.3. Immunoblotting Cell extracts were prepared in a buffer consisting of 50 mM Tris–HCl, 5 mM EDTA, 5 mM EGTA, 20 mg/ml leupeptin, 20 mg/ml aprotinin, 100 mg/ml PMSF, and 50 mM DTT. For each sample, the protein content was determined using the Bio-Rad protein assay. Samples (20 mg protein/ lane) of cell extracts were run on 5% SDS-polyacrylamide gels, transferred to nitrocellulose membranes (0.45 mm; Schleicher and Schuell, Keene, NH), and probed with a mouse monoclonal antibody to dystrophin (Mandys 8; Sigma Chemical, St. Louis, MO) at a 1:400 dilution. The specific antibody reaction was detected by an enhanced chemiluminescence system (Amersham Corp., Arlington Heights, IL). 2.4. Immunocytochemistry Myotube cultures were fixed in 2% formaldehyde in PBS for 5 min at room temperature followed by 100% methanol at −20°C for 5 min. The cells were then rinsed in PBS. All further incubations were carried out at room temperature and all rinses and dilutions were with a blocking solution consisting of 5% horse serum and 0.5% Triton-X 100 in PBS. An initial blocking step was performed with this solution for 30 min. A mouse monoclonal antibody to dystrophin (Dys1 - Novocastra Laboratories; Vector Laboratories, Burlingame, CA) was applied neat for 30 min. After rinsing, a Texas Red-coupled secondary antibody (Cappel Research Products, Durham, NC) was applied for 30 min at a dilution of 1:250. The cells were examined and photographed with a Zeiss Axioskop microscope. 2.5. Cell survival assay Myoblasts were plated in GM into 24 well plates at a density of 7.5 × 104 cells/well. After 1 day, the medium was changed to DM. Cells were differentiated for 4–5 days (myotube formation began after day 2), and cellular toxins were added to each well for 12–18 h as indicated. Cell survival was determined using a kit for measuring lactate dehydrogenase (LDH) activity according to the instructions of the manufacturer (Promega, Madison, WI). Briefly,
16
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
cellular LDH was used as a measure of the number of surviving cells in each well. After the incubation period, the supernatant was removed, the remaining cells were rinsed, and the cellular LDH activity was measured. Cell death was calculated as a percentage, with 0% cell death being the value of cellular LDH in wells with no added toxin, and 100% cell death being the value of cellular LDH remaining after addition of 0.9% SDS to the culture medium just prior to the end of the incubation period. Thus, for each cell population, there were internal positive and negative controls for the assay itself. All data are from assays done in triplicate. During initial studies to validate the use of the LDH assay, cell death was corroborated in parallel myoblast cultures by hemacytometer counts of total and viable cell numbers using trypan blue exclusion after trypsinization, and there was an excellent agreement between the two methods. It was also routine to estimate cell death by visual inspection of the cultures at the end of the incubation period as a further corroboration (estimating to the closest 10% over the range of 0% (untreated cultures) to 100% cell death), and these estimates paralleled the LDH results as well. For myotube cultures, hemacytometer counts are not accurate, so corroboration of LDH assay results was always done by visual inspection of the cultures before the assay. There was never a discordance between the general level of cell death by the two methods, but of course estimates of cell death by visual inspection were less accurate than those by the LDH assay. 2.6. Cellular toxins Cellular toxins were divided into ‘pro-oxidants’ (those whose mechanism of toxicity was primarily by promoting oxidative injury) and ‘non-oxidants’ (those whose mechanism of injury was not primarily due to free radical injury). In each case, dose-response curves were done and the levels of cell death were determined over time. The incubation time chosen for these studies was in the range of 12–18 h because this resulted in a ‘steady-state’ level of cell death in response to the one-time addition of each toxin to the medium across the concentration ranges listed below. All reagents were purchased from Sigma Chemical (St. Louis, MO) unless otherwise indicated. Pro-oxidants tested were: H2O2: 0.01–31.6 mM; H2O2 generates hydroxyl radicals by metal-catalyzed reactions (Fenton chemistry) [21]. Paraquat: 0.01–31.6 mM; paraquat generates superoxide intracellularly [22]. Menadione: 0.1 mM–1 mM; menadione is a quinone that generates free radicals intracellularly by undergoing repeated redox cycles [23]. AAPH (2,2′-amino azo bis amidopropane hydrochloride; Wako Chemicals, Richmond, VA): 0.1–100 mM;
AAPH generates peroxyl radicals that can initiate lipid peroxidation [24]. SNP (sodium nitroprusside): 0.01–31.6 mM; SNP generates nitric oxide [25] which combines with endogenous superoxide to form the potent reactive oxygen species, peroxynitrite [26]. Non-oxidants tested were: Staurosporine: 1 nM–1 mM; staurosporine has been shown to promote apoptotic cell death in a variety of cell types by the inhibition of protein kinases [27]. A23187: 0.1–100 mg/ml; the primary mechanism of toxicity is via rapid rises in intracellular calcium and activation of proteases, phospholipases, and endonucleases [28]. CCCP (carbonyl cyanide m-chlorophenylhydrazone): 0.1–100 mM; CCCP is an inhibitor of mitochondrial respiration. The primary mechanism of toxicity of mitochondrial inhibitors is via reduction of cellular ATP levels, and the resulting cellular injury is independent of oxidant injury [29]. 2.7. Statistical analysis Comparisons between samples were done by analysis of variance (ANOVA). Differences were considered statistically significant at the P , 0.05 level.
3. Results Undifferentiated skeletal myoblasts were derived from neonatal mice [20] of both the C57 and the mdx strains. Myoblasts from C57 and mdx muscle behaved similarly in assays of cell growth (population doubling times were 19.4 ± 0.8 h for C57 myoblasts, and 18.9 ± 0.5 h for mdx myoblasts) and differentiation (the fusion indices were 0.38 ± 0.04 in C57 cultures and 0.39 ± 0.02 in mdx cultures after 3 days in DM). When grown in mitogen-poor medium, skeletal myoblasts differentiated into multinucleated myotubes (Fig. 1) which expressed many of the morphological and biochemical properties of mature muscle cells in vivo (e.g., expression of myosin heavy chain, skeletal muscle actin, creatine phosphokinase), except for the fact that mdx myotubes did not express dystrophin (Fig. 1). We exposed both C57 and mdx myotube cultures to prooxidant compounds to compare their susceptibilities to oxidative stress. Fig. 2 shows dose-response curves of cell death as a function of concentration of two compounds that induce oxidative stress, the reactive oxygen species H2O2 and the superoxide-generating compound, paraquat. For both compounds, mdx myotubes were more susceptible to the oxidative injury than were normal myotubes as evidenced by greater cell death across a wide concentration range. In order to compare the susceptibility of the two
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
17
Fig. 1. Differentiation of C57 and mdx muscle cells in culture. (A) Myotube formation by C57 and mdx muscle cells. Myoblast cultures derived from C57 and mdx mice were induced to differentiate by culturing in low serum medium. After 5 days, myoblast elongation and fusion lead to multinucleated myotube formation in each culture. The top panels show phase contrast images of the myotube cultures. The bottom panels show immunofluorescent staining for dystrophin in the two populations. Dystrophin expression was detected only in the C57 myotubes. (B) Time course of expression of dystrophin in myotube cultures. Cultures of muscle cells from both C57 and mdx mice were maintained in DM, and each day cultures were analyzed by immunoblot analysis for the expression of dystrophin. Dystrophin expression increased during the first several days in C57 cultures, and plateaued after 3–4 days. No dystrophin expression was observed in the mdx cultures.
Fig. 2. Dystrophin-deficient myotubes are more susceptible to oxidative injury than are normal myotubes. Both C57 and mdx myotube cultures were subjected to oxidative stress by the addition of the pro-oxidant compounds (A) H2O2 or (B) paraquat to the culture medium. Sixteen hours after exposure to different concentrations of the pro-oxidants, the extent of cell death was determined by the LDH assay. In both cases, mdx myotubes exhibited greater sensitivity to oxidative stress than did C57 myotubes. Data are expressed as mean ± SD (n = 4).
18
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
Fig. 3. Comparisons of susceptibilities of C57 and mdx muscle cells to oxidative stress and non-oxidative metabolic stresses. Experiments were done as in Fig. 2, and the extent of cell death was compared at concentrations of the toxins that resulted in sub-maximal cell death. Data are expressed as mean ± SD (n = 4). (A) Susceptibility of C57 and mdx myotubes to oxidative stress. The pro-oxidants (see Section 2) were tested at the following concentrations: 0.316 mM H2O2; 5 mM menadione; 2 mM paraquat; 16.5 mM AAPH; 1 mM SNP. Despite differences in mechanisms of producing oxidative stress among the different toxins, mdx myotubes were more susceptible than C57 myotubes to all of the compounds tested. (B) Susceptibility of C57 and mdx myotubes to non-oxidant toxins (see Section 2). The non-oxidants were tested at the following concentrations: 0.01 mM staurosporine; 2 mg/ml A23187; 10 mM CCCP. The two populations of myotubes were equally susceptible to all of the non-oxidant compounds tested.
cell populations, we chose concentrations of the toxins that produced sub-maximal levels of cell death in mdx cultures and detectable levels of cell death in C57 cultures. Fig. 3A summarizes the results of these and similar experiments with a variety of pro-oxidant compounds, each of which induces oxidative stress by a different mechanism (see Section 2). In each case, the susceptibility of dystrophin-deficient myotubes to oxidative injury was significantly greater than that of the normal myotubes. Although these results demonstrated that dystrophin-deficient muscle was more sensitive than normal muscle to oxidative stress, we subjected the cells to non-oxidant forms of injury to test the possibility that dystrophin deficient muscle was just more susceptible to any form of metabolic stress. Thus, C57 and mdx myotube cultures were exposed to toxins whose mechanisms of injury are not via free radical mechanisms (see Section 2). C57 and mdx myotube cultures were equally sensitive to the toxic effects of
these agents at each concentration tested, and the effects of the dose of each toxin that produced approximately 50% cell death is shown (Fig. 3B). Certainly, any significant difference in susceptibility of the cells to these toxins would be apparent at these intermediate concentrations. These results suggest that dystrophin-deficient muscle cells are specifically more susceptible to oxidative stress, and not simply more susceptible to any form of metabolic stress that might cause cellular injury. C57 and mdx mice are from the same genetic background, and differ genetically by the presence of a mutation in the dystrophin gene in the mdx strain [30]. To confirm that the difference in susceptibility to oxidative stress of the two strains was indeed due to the difference in dystrophin expression, we compared the susceptibility of undifferentiated myoblasts from the two strains to oxidative stress. Since dystrophin is not expressed in myoblasts, the two populations should be identical biochemically, and we predicted that they would be equally susceptible to any form of injury. Indeed, Fig. 4 shows that the C57 and mdx myoblasts were equally susceptible to the toxic effects of paraquat. There was no difference in susceptibility to every metabolic toxin tested, including the other pro-oxidants H2O2 and menadione, as well as the control toxins staurosporine and A23187. Finally, as a further indication that dystrophin expression confers a certain level of resistance to oxidative injury, we compared the susceptibilities of myotubes from four normal strains of mice: C57, C3H, SJL, and BALB/c. All four strains express dystrophin and have no intrinsic muscle disorder. Among the four strains tested, there were no differences in their susceptibilities to pro-oxidants (Fig. 5), nor was there any difference in susceptibilities to non-oxidants. This was true over a range of concentrations that resulted in anywhere from 25% to 75% cell death, the most sensitive
Fig. 4. Susceptibilities of C57 and mdx myoblasts to oxidative stress. Undifferentiated myoblasts were maintained in growth medium and exposed to different concentrations of paraquat for 18 h. Cell death was measured using the LDH assay. Data are presented as mean ± SD (n = 5). Unlike myotube cultures, mdx myoblasts were not more susceptible to oxidative stress than were C57 myoblasts.
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
Fig. 5. Susceptibilities of myotubes from different normal strains of mice to oxidative stress. In addition to the C57 strain, which is the parental strain of the mdx, we tested the relative susceptibilities of myotube cultures from different dystrophin-positive strains to oxidative stress. These data show the results of the addition of either (A) menadione (10 mM) or (B) paraquat (5 mM) to the culture medium of myotube cultures from the C57, C3H, BALB/c, and SJL strains of mice. Cell death was measured after 12 h of exposure, and data are presented as mean ± SD (n = 4). There was no significant difference between the results of any of the strains; all were equally susceptible to oxidative stress.
range for detecting any differences between the cell populations. Thus, among five independently derived primary muscle cell cultures, only the dystrophin-deficient myotubes showed an increased susceptibility to oxidative stress.
4. Discussion Selective muscle involvement is a hallmark of the muscular dystrophies, indicating that genetic defects present in all muscles do not affect all muscles equally [1,2,5]. In muscular dystrophies due to dystrophin deficiency, there are examples of specific muscle sparing in humans and
19
other animal species. In particular, in the mdx mouse there is no evidence of muscle necrosis for the first several weeks postnatally [9]. There follows a period of widespread necrosis over the next 6–8 weeks, after which there is minimal necrosis in limb muscles for the remainder of the adult life. By contrast, there is also evidence in all species examined that certain muscles are severely affected [1,2,5]. Clearly, epigenetic mechanisms play an important role in determining whether a muscle will succumb to the degenerative process, when the degeneration will begin (or end), and how severely a muscle will be affected. The relative susceptibility or resistance of each muscle will account for phenotypic variability. Examining cellular susceptibility in vivo is particularly complex because of the many uncontrolled variables that may affect cell survival or cell death. For this reason, cellular susceptibility is commonly studied in culture where pharmacologic agents are used to mimic physiologic stresses, such as oxidative stress, that may occur in vivo [31–33]. We have taken this approach by using a purified population of myogenic cells. While lacking the complexities of mature muscle tissue, these studies allow analysis using a single end-point that occurs both in vivo and in vitro, namely cell death. Our results suggest that dystrophin deficiency renders muscle susceptible to oxidative stress, but not merely to any form of metabolic stress. The fact that both extreme exercise [12,13] and muscle ischemia [10,11] can generate lesions that resemble dystrophic changes, and that such lesions are mediated by oxidative injury [34,35], supports the hypothesis that a muscular dystrophy could be a consequence of an increased susceptibility to this form of metabolic stress. Furthermore, similarities between the pathology of muscular dystrophies and the myopathy of vitamin E deficiency, a condition of increased susceptibility to oxidative stress, have long been noted [14]. Recently, we have also documented a muscular dystrophy caused by a genetic alteration in free radical metabolism. In a transgenic mouse that overexpresses the enzyme Cu,Zn-superoxide dismutase [36], we have found a profound muscular dystrophy that begins between 2 and 4 months of age (unpublished observations). Overexpression of this enzyme has been reported to sensitize cells to oxidative stress by producing elevated levels of H2O2 [33,37]. As a model for a muscular dystrophy specifically due to oxidative stress, this mouse strain should prove to be very informative. The role of free radical mediated injury in the pathogenesis of the muscular dystrophies has been extensively studied in vivo [14]. Working with mammalian and avian dystrophic muscle, investigators have demonstrated increased lipid peroxidation [14,38], increased oxidation of proteins [16,17], and induction of antioxidant enzymes [39,40]. We have studied the role of oxidative injury in the mdx mouse in vivo by focusing on the pre-necrotic state, since the onset of muscle degeneration does not begin until about 3 weeks of age. We have found an increase in lipid
20
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21
peroxidation in mdx muscle preceding the onset of muscle necrosis [19]. The free radical hypothesis complements many current ideas on the pathogenesis of the disease [3]. The ‘mechanical damage hypothesis’, the ‘leaky membrane hypothesis’, and the ‘calcium hypothesis’ all address some of the cardinal features of dystrophic muscle, there is evidence to support aspects of each of these theories [41–43], and they are not mutually exclusive. Certainly, there appears ultimately to be a disruption of membrane integrity with subsequent calcium influx leading to cell necrosis, but the biochemical process that initiates this chain of events is unspecified and the resistance of certain muscles to the degenerative process remains unexplained. The free radical hypothesis suggests just such a biochemical cascade: normal cellular metabolism produces free radicals, free radicals cause lipid peroxidation, lipid peroxidation causes membrane breakdown and cell necrosis. According to the hypothesis and as suggested by our data, the absence of dystrophin would be associated with an increased susceptibility to oxidative injury, whereas the membrane of normal muscle would be more resistant. Cellular antioxidant capacity differs among muscles and as a function of age [44,45], and such changes in cell protective mechanisms may explain how a disease resulting from a constant genetic defect could be manifested differently over space and time [18]. As dystrophin is not known to have any antioxidant properties, the question arises as to how dystrophin may function to protect the cell against free radical injury, and, conversely, how the absence of dystrophy could render a cell susceptible to free radical injury. The recent demonstration of the association of the dystrophin complex with an enzyme, namely nitric oxide synthase (NOS), involved in free radical metabolism is intriguing [46,47]. This is the first demonstration of the association of dystrophin with a cellular metabolic pathway, and that pathway happens to involve free radical metabolism, namely the generation of the free radical nitric oxide (NO). How this association, and its disruption in dystrophin-deficient muscle [46,47], may relate to the pathogenesis of muscular dystrophies remains to be determined. Alterations in NOS activity could alter NO levels in the cell, and NO has been shown to have both protective and destructive actions in cellular oxidative injury [48]. We are currently exploring the role of NO in muscle cell oxidative metabolism and free radical injury. The possibility that oxidative stress may play an important role in the pathogenesis of muscle degeneration in the dystrophies could have profound therapeutic consequences. Clinical trials of antioxidants have been discouraging in Duchenne muscular dystrophy [49,50], but these have always involved patients long after the onset and progression of the degeneration. Our results suggest that oxidative injury might contribute to the very first pathologic change, namely cell death, and that antioxidant therapy could be effective only if started early enough to prevent muscle cell necrosis. The mdx mouse is an ideal model system for
such trials, since the onset of muscle necrosis occurs after approximately 3 weeks of age [3]. The benefit of early intervention with antioxidant therapy could be tested in the mdx mouse to determine if such trials would be warranted in humans with Duchenne muscular dystrophy.
Acknowledgements The authors would like to thank Dr. Grace Pavlath of Emory University for supplying myoblast cultures from BALB/c and SJL strains. This work was supported in part by a Howard Hughes Summer Fellowship from the Department of Biological Sciences at Stanford University to AAF and by grants from the Department of Veterans Affairs, the Muscular Dystrophy Association, and the American Academy of Neurology Education and Research Foundation to TAR. References [1] Emery AEH. Duchenne muscular dystrophy. New York: Oxford University Press, 1993. [2] Hoffman EP, Gorospe JRM. The animal models of Duchenne muscular dystrophy: windows on the pathophysiological consequences of dystrophin deficiency. Curr Top Memb 1991;38:113–154. [3] McArdle A, Edwards RHT, Jackson MJ. How does dystrophin deficiency lead to muscle degeneration? – Evidence from the mdx mouse. Neuromusc Disord 1995;5:445–456. [4] Valentine BA, Cooper BJ. Canine X-linked muscular dystrophy: selective involvement of muscles in neonatal dogs. Neuromusc Disord 1991;1:31–38. [5] Stedman HH, Sweeney HL, Shrager JB. et al. The mdx mouse diaphragm reproduces the degenerative changes of Duchenne muscular dystrophy. Nature 1991;352:536–538. [6] Boland B, Himpens B, Denef JF, Gillis JM. Site-dependent pathological differences in smooth muscles and skeletal muscles of the adult mdx mouse. Muscle Nerve 1995;18:649–657. [7] Minetti C, Ricci E, Bonilla E. Progressive depletion of fast alphaactinin-positive muscle fibers in Duchenne muscular dystrophy. Neurology 1991;41:1977–1981. [8] Karpati G, Carpenter S, Prescott S. Small-caliber skeletal muscle fibers do not suffer necrosis in mdx mouse dystrophy. Muscle Nerve 1988;11:795–803. [9] Tanabe Y, Esaki K, Nomura T. Skeletal muscle pathology in X chromosome-linked muscular dystrophy (mdx) mouse. Acta Neuropathol 1986;69:91–95. [10] Mendell JR, Engel WK, Derrer EC. Duchenne muscular dystrophy: functional ischemia reproduces its characteristic lesions. Science 1971;172:1143–1145. [11] Engel WK. Duchenne muscular dystrophy: a histologically based ischemia hypothesis and comparison with experimental ischemia myopathy. In: Pearson CM, Mostofi FK, editors. The striated muscle. Baltimore: Williams and Wilkins, 1973:453–472. [12] Vihko V, Rantamaki J, Salminen A. Exhaustive physical exercise and acid hydrolase activity in mouse skeletal muscle. Histochemistry 1978;57:237–249. [13] Irintchev A, Wernig A. Muscle damage and repair in voluntarily running mice: strain and muscle differences. Cell Tissue Res 1987;249:509–521. [14] Murphy ME, Kehrer JP. Oxidative stress and muscular dystrophy. Chem-Biol Interact 1989;69:101–178.
T.A. Rando et al. / Neuromuscular Disorders 8 (1998) 14–21 [15] Brown RH. Free radicals, programmed cell death, and muscular dystrophy. Curr Opin Neurol 1995;8:373–378. [16] Hauser E, Hoger H, Bittner R, Widhalm K, Herkner K, Lubec G. Oxyradical damage and mitochondrial enzyme activities in the mdx mouse. Neuropediatrics 1995;26:260–262. [17] Haycock JW, MacNeil S, Jones P, Harris JB, Mantle D. Oxidative damage to muscle protein in Duchenne muscular dystrophy. NeuroReport 1996;8:357–361. [18] Ragusa FJ, Chow CK, St Clair DK, Porter JD. Extraocular, limb, and diaphragm muscle group-specific antioxidant enzyme activity patterns in control and mdx mice. J Neurol Sci 1996;139:180–186. [19] Rando TA, Disatnik M-H, Dhawan J, Yu Y, Whirl MM. Oxidative injury precedes muscle cell necrosis in dystrophin-deficient (mdx) mice. Neurology 1997;48:A442. [20] Rando TA, Blau HM. Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol 1994;125:1275–1287. [21] Lloyd RV, Hanna PM, Mason RP. The origin of the hydroxyl radical oxygen in the Fenton reaction. Free Rad Biol Med 1997;22:885–888. [22] Krall J, Bagley AC, Mullenbach GT, Hallewell RA, Lynch RE. Superoxide mediates the toxicity of paraquat for cultured mammalian cells. J Biol Chem 1988;263:1910–1914. [23] Thor H, Smith MT, Hatzell P, Bellomo G, Jewell SA, Orrenius S. The metabolism of menadione (2-methyl-1,4-naphthoquinone) by isolated hepatocytes. A study of the implications of oxidative stress in intact cells. J Biol Chem 1982;257:12419–12425. [24] Dooley MM, Sano N, Kawashima H, Nakamura T. Effects of 2,2′azobis (2-amidinopropane) hydrochloride in vivo and protection by vitamin E. Free Rad Biol Med 1990;9:199–204. [25] Feelisch M, Noack EA. Correlation between nitric oxide formation during degradation of organic nitrates and activation of guanylate cyclase. Eur J Pharmacol 1987;139:19–30. [26] Beckman JS, Beckman TW, Chen J, Marshall PA, Freeman BA. Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide. Proc Natl Acad Sci USA 1990;87:1620–1624. [27] Jacobson MD, Raff MC. Programmed cell death and Bcl-2 protection in very low oxygen. Nature 1995;374:814–816. [28] Nicotera P, Bellomo G, Orrenius S. Calcium-mediated mechanisms in chemically induced cell death. Annu Rev Pharmacol Toxicol 1992;32:449–470. [29] Weinberg JM, Davis JA, Abarzua M, Kaini T, Kunkel R. Protection by glycine of proximal tubules from injury due to inhibitors of mitochondrial ATP production. Am J Physiol 1990;258:C1127–C1140. [30] Sicinski P, Geng Y, Ryder-Cook AS, Barnard EA, Darlison MG, Barnard PJ. The molecular basis of muscular dystrophy in the mdx mouse: a point mutation. Science 1989;244:1578–1580. [31] Zhong L-T, Sarafian T, Kane DJ. et al. bcl-2 Inhibits death of central neural cells induced by multiple agents. Proc Natl Acad Sci USA 1993;90:4533–4537. [32] Bonfoco E, Krainc D, Ankarcrona M, Nicotera P, Lipton SA. Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-d-aspartate or nitric oxide/superoxide in cortical cell cultures. Proc Natl Acad Sci USA 1995;92:7162–7166.
21
[33] Amstad P, Moret R, Cerutti P. Glutathione peroxidase compensates for the hypersensitivity of Cu,Zn-superoxide dismutase overproducers to oxidant stress. J Biol Chem 1994;269:1606–1609. [34] Radak Z, Asano K, Inoue M. et al. Superoxide dismutase derivative reduces oxidative damage in skeletal muscle of rats during exhaustive exercise. J Appl Physiol 1995;79:129–135. [35] Fantini GA, Yoshioka T. Deferoxamine prevents lipid peroxidation and attenuates reoxygenation injury in postischemic skeletal muscle. Am J Physiol 1993;264:H1953–H1959. [36] Epstein CJ, Avraham KB, Lovett M. et al. Transgenic mice with increased Cu/Zn-superoxide dismutase activity: animal model of dosage effects in Down syndrome. Proc Natl Acad Sci USA 1987;84:8044–8048. [37] Amstad P, Peskin A, Shah G. et al. The balance between Cu,Znsuperoxide dismutase and catalase affects the sensitivity of mouse epidermal cells to oxidative stress. Biochem 1991;30:9305–9313. [38] Mizuno Y. Changes in superoxide dismutase, catalase, glutathione peroxidase, and glutathione reductase activities and thiobarbituric acid-reactive product levels in early stages of development in dystrophic chickens. Exp Neurol 1984;84:58–73. [39] Burr IM, Asayama K, Fenichel G. Superoxide dismutase, glutathione peroxidase, and catalase in neuromuscular disease. Muscle Nerve 1987;10:150–154. [40] Austin L, de Niese M, McGregor A, Arthur H, Gurushinghe A, Gould MK. Potential oxyradical damage and energy status in individual muscle fibres from degenerating muscle diseases. Neuromusc Disord 1992;2:27–33. [41] Petrof BJ, Shrager JB, Stedman HH, Kelly AM, Sweeney HL. Dystrophin protects the sarcolemma from stresses developed during muscle contraction. Proc Natl Acad Sci USA 1993;90:3710–3714. [42] Turner PR, Fong P, Denetclaw WF, Steinhardt RA. Increased calcium influx in dystrophic muscle. J Cell Biol 1991;115:1701–1712. [43] Rowland LP. Biochemistry of muscle membranes in Duchenne muscular dystrophy. Muscle Nerve 1980;3:3–20. [44] Powers SK, Criswell D, Lawler J. et al. Influence of exercise and fiber type on antioxidant enzyme activity in rat skeletal muscle. Am J Physiol 1994;266:R375–R380. [45] Oh-Ishi S, Kizaki T, Yamashita H. et al. Alterations of superoxide dismutase iso-enzyme activity, content, and mRNA expression with aging in rat skeletal muscle. Mech Ageing Dev 1995;84:65–76. [46] Brenman JE, Chao DS, Xia H, Aldape K, Bredt DS. Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell 1995;82:743– 752. [47] Chang W-J, Iannaccone ST, Lau KS. et al. Neuronal nitric oxide synthase and dystrophin-deficient muscular dystrophy. Proc Natl Acad Sci USA 1996;93:9142–9147. [48] Darley-Usmar V, Wiseman H, Halliwell B. Nitric oxide and oxygen radicals: a question of balance. FEBS Lett 1995;369:131–135. [49] Roelofs RI, de Arango GS, Law PK, Kinsman D, Buchanan DC, Park JH. Treatment of Duchenne’s muscular dystrophy with penicillamine. Arch Neurol 1979;36:266–268. [50] Fenichel GM, Brooke MH, Griggs RC. et al. Clinical investigation in Duchenne muscular dystrophy: penicillamine and vitamin E. Muscle Nerve 1988;11:1154–1168.