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Published online before print September 6, 2007
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From the Basic and Clinical Myology Laboratory, Department of Physiology, The University of Melbourne, Melbourne, Victoria, Australia
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Abstract |
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The muscular dystrophies define a group of genetic disorders characterized by progressive skeletal muscle wasting and weakness. The most severe and rapidly progressing of these conditions is Duchenne muscular dystrophy, affecting 
1 in 3500 males born worldwide.2
The absence of a cytoskeletal protein, dystrophin, renders dystrophic muscles extremely fragile and easily damaged by everyday activities, especially by contractions where muscles are activated and then stretched forcibly.3,4
This increased susceptibility to contraction-mediated injury results in ongoing cycles of muscle fiber degeneration and less than adequate regeneration and contributes to an etiology of progressive muscle wasting and weakness.5
Although animal models can reproduce some (or most) of the pathologies associated with muscular dystrophies, models of muscle injury such as crush or strain injuries can be problematic especially for generating reproducible responses in animal models. On the other hand, injecting muscles with a myotoxic agent such as notexin or cardiotoxin results in complete and reproducible muscle fiber degeneration6 and the ability to monitor the events of and mechanisms controlling muscle fiber regeneration.
Regardless of the initial injury, effective muscle regeneration is dependent on the timed induction of myogenic regulatory factors and growth factors, including insulin-like growth factor-I (IGF-I).1,7 IGF-I activates both myoblast proliferation and subsequently differentiation, two processes that are crucial to muscle repair and regeneration.8,9 The importance of IGF-I in skeletal muscle regeneration has been demonstrated in transgenic mice, where overexpression of IGF-I localized to skeletal muscle maintained regenerative capacity in aged mice10 and reduced the dystrophic pathology in mdx mice, an animal model of Duchenne muscular dystrophy.11,12 In addition, we have shown previously that exogenous administration of recombinant human IGF-I (rhIGF-I) increased the rate of functional recovery after myotoxic injury13 and improved the dystrophic pathology in mdx mice.14-16 Although rhIGF-I administration and transgenic IGF-I overexpression have beneficial effects on skeletal muscle, their mechanism of action seems to be vastly different. For example in mice, transgenic IGF-I overexpression resulted in marked muscle hypertrophy,11 an effect not observed following rhIGF-I administration.14-16 This discrepancy is likely due to different effects of IGF-binding proteins (IGFBPs) on IGF-I in the bloodstream (systemic delivery) compared with muscle-specific transgenic overexpression of IGF-I.
The biological actions of IGF-I in vivo are strongly mediated by IGFBPs, which bind 99% of IGF-I in the circulation.17
This large reservoir of bound, biologically inactive IGF-I has a prolonged half-life and is protected from degradation.17-19
IGFBPs are believed to primarily transport IGF-I to target tissues, modulate IGF-I action, and prevent hypoglycemia.17-19
In general, IGFBPs are thought to inhibit IGF-I actions in vitro and in vivo by preventing IGF-I binding to cell surface receptors.20-22
Although the effects of rhIGF-I administration and IGF-I overexpression on skeletal muscle regeneration have been well characterized,10-12,14-16 the role of IGFBPs in skeletal muscle regeneration remains poorly understood. Thus, the aim of the current study was to examine the effects of IGFBP inhibition on muscle regeneration after myotoxic injury and also in mdx dystrophic mice. We tested the hypothesis that inhibition of endogenous IGFBPs would enhance skeletal muscle regeneration and improve the dystrophic pathology in a manner similar to exogenous rhIGF-I administration.
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Materials and Methods |
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All procedures were approved by the Animal Experimentation Ethics Committee of The University of Melbourne and conformed to the Guidelines for the Care and Use of Experimental Animals as described by the National Health and Medical Research Council of Australia (Canberra, ACT, Australia). To examine the effect of IGFBP inhibition on muscle regeneration, male C57BL/10ScSn (BL/10) mice (12 to 14 weeks old, n > 5 per group) were used. To determine the role of IGFBPs in dystrophic skeletal muscles, male C57BL/10ScSn-mdx/J dystrophic (mdx) mice (8 to 10 weeks old, n = 8 per group) were used. All mice were obtained from the Animal Resource Centre (Canning Vale, WA, Australia) and housed in the Biological Research Facility at The University of Melbourne under a 12-hour light/dark cycle, with drinking water and standard chow provided ad libitum. For both experiments treated mice received continuous subcutaneous (s.c.) administration of the IGF-I aptamer NBI-31772 (6 mg/kg per day in 50% dimethyl sulfoxide and 50% polyethylene glycol), whereas control mice received vehicle alone (50% dimethyl sulfoxide and 50% polyethylene glycol). Continuous systemic delivery of NBI-31772 (or vehicle alone) was achieved by subcutaneous implantation of a micro-osmotic pump (Alzet model 1002; Alzet, Cupertino, CA) as described previously.14 NBI-31772 is a nonpeptide ligand that has high-affinity binding for all six IGF binding proteins,23 and in vitro binding assays have revealed that NBI-31772 displaces IGF-I from its interaction with binding proteins, thus releasing "free" biologically active IGF-I.24 BL/10 mice received NBI-31772 infusion for a period of 10, 14, or 21 days, whereas mdx mice received NBI-31772 for 28 days.
Myotoxic Injury of Tibialis Anterior Muscles
In BL/10 mice, tibialis anterior (TA) muscles were injured using the myotoxic agent notexin, as described previously.13 In brief, mice were anesthetized deeply with sodium pentobarbitone (60 mg/kg i.p., Nembutal; Rhone Merieux, Pinkenba, QLD, Australia), and an adequate depth of anesthesia was maintained, such that the mice were unresponsive to tactile stimulation. The right hindlimb was shaved, and a small portion of the anterior aspect of the TA muscle was surgically exposed. The muscle was injected with 40 µl of notexin (10 µg/ml in isotonic saline; Latoxan, Valence, France) with a 29-gauge fixed needle. Care was taken to prevent leakage of notexin from the muscle. An injection volume of 40 µl was the maximum holding capacity of a TA muscle and reliably produced complete degeneration of all myofibers.6,13 After the intramuscular injection, the skin incision was closed with surgical (Michel) clips (Aesculap, Tuttlingen, Germany), and the mouse was allowed to recover on a heat pad.
Contractile Properties Measured In Situ
The methods for in situ contractile analysis of TA muscles from mice have been described in detail elsewhere.16 In brief, TA muscles from injured and control BL/10 mice were stimulated by supramaximal (10 V) 0.2-ms square wave pulses of 300 ms in duration delivered via two wire electrodes adjacent to the femoral nerve. Optimum muscle length (Lo) was determined from maximum isometric twitch force (Pt), and maximum isometric tetanic force (Po) was recorded from the plateau of the frequency-force relationship.
Contractile Properties Measured In Vitro
The contractile properties of extensor digitorum longus (EDL) and soleus muscles and of diaphragm muscle strips from mdx mice were analyzed in vitro, as described previously.25 In brief, EDL, soleus, and diaphragm muscle preparations were stimulated by supramaximal (40 V) 0.2-ms square wave pulses of 350-, 1200-, and 450-ms duration, respectively, delivered via platinum plate electrodes that flanked both sides of the muscle. Lo was determined at Pt and Po was recorded from the plateau of the frequency-force curve. The EDL and soleus muscles from the right hindlimb, as well as a diaphragm muscle strip, were assessed for their susceptibility to contraction-induced injury, whereas the EDL and soleus muscles from the left hindlimb, as well as another diaphragm muscle strip, were assessed for their resistance to a fatiguing stimulation protocol.
The contraction-induced injury protocol used in this study was similar to others described previously.26,27 Muscles were lengthened at a velocity of two Lf/s at progressively increasing magnitudes of stretch beyond Lf (5, 10, 20, 30, 40, and 50%), with maximum isometric force being determined after each lengthening contraction. The "force deficit" after contraction-induced damage was determined by calculating the difference between the Po measured 2 minutes after the lengthening contractions and Po determined before any of the lengthening contractions and is expressed as a percentage of the maximum Po determined before the protocol of lengthening contractions.16,26
Muscle fatigue was assessed using a standard fatigue stimulation protocol we have described previously.25 Muscles were stimulated maximally once every 4 seconds for 4 minutes, with maximum force recorded every minute. During recovery, maximum force was determined at 5, 10, and 15 minutes after completion of the fatigue protocol.
Skeletal Muscle Morphology
At the completion of all of the contractile function analyses, the muscles were mounted in embedding medium, frozen in thawing isopentane, and stored at –80°C. A portion of each muscle was cryosectioned transversely (8 µm) through the midbelly region. Muscle sections were stained with hematoxylin and eosin (H&E) to determine general muscle architecture and to determine the cross-sectional area (CSA) of individual muscle fibers. Median values for CSA were calculated from at least 200 individual muscle fibers per cross section, as described previously.16
Myosin Heavy Chain Analysis
Myosin heavy chain (MyHC) isoform composition within TA muscles of BL/10 mice was determined via sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Muscle samples were homogenized on ice in phosphate-buffered saline and centrifuged at 300 x g for 2 minutes at 4°C. The pellet was resuspended in 100 µl of Guba-Straub buffer (300 mmol/L NaCl, 100 mmol/L NaH2PO4H2O, 50 mmol/L NaH2PO4, 1 mmol/L MgCl2·6H2O, 10 mmol/L Na2H2P2O7, and 10 mmol/L ethylenediamine tetraacetic acid) and centrifuged at 12,000 x g for 15 minutes at 4°C. The supernatant was diluted with 1.5 ml of H2O and refrigerated overnight. The following day, the supernatant was centrifuged again at 12,000 x g for 15 minutes at 4°C, and the pellet was resuspended in 300 µl of Guba-Straub buffer containing 10% glycerol. Protein concentration was determined via the method of Bradford,28 and all samples were equalized. Ten micrograms of protein was added to 90 µl of Guba-Straub buffer and 30 µl of bromphenol blue solution. Immediately before gel loading, samples were heated at 95°C for 5 minutes, after which 4 µl of sample was loaded per well. MyHC isoforms were separated on an 11% acrylamide-bis (50:1) gel by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The gels were run at 70 V at 4°C for 36 hours. Protein bands were visualized using the InVitrogen Silver Stain kit (LC6100; Victoria, Australia). The gels were scanned and band densities analyzed using the image analysis software described previously.29
Citrate Synthase Activity
Citrate synthase activity was determined in homogenates prepared from EDL, soleus, and diaphragm muscles using a citrate synthase assay kit (CS0720; Sigma-Aldrich, St. Louis, MO).30 Total muscle protein was determined in triplicate by the method of Bradford,28 and the protein concentration of all samples was equalized. Citrate synthase activity was determined in triplicate based on the formation of 2-nitro-5-thiobenzoic acid at a wavelength of 412 nm at 25°C on a spectrophotometer (Multiskan Spectrum; Thermo Electron Corporation, Waltham, MA). In each well, 8 µl of sample was added to a reaction medium containing 178 µl of assay buffer, 2 µl of 30 mmol/L acetyl coenzyme A, and 10 mmol/L 2-nitro-5-thiobenzoic acid. The baseline solution absorbance was recorded, reactions were initiated by the addition of 10 µl of oxaloacetic acid, and the change in absorbance measured every 15 seconds for 2 minutes.
Statistical Analysis
All values are expressed as mean ± SEM unless specified otherwise. Groups were compared using two-way analysis of variance where appropriate. The level of significance was set at P < 0.05 for all comparisons. Bonferroni’s post hoc MCP was used to determine significant differences between groups. CSA data were analyzed using the Anderson-Darling Normality Test. Muscle fiber CSA was not normally distributed, and so the 95% confidence interval of the median was used and differences were considered significant when no overlap existed between the 95% confidence interval of the median.
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Results |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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Administration of NBI-31772 did not alter the mass of regenerating TA muscles at 10, 14, or 21 days after injury (Table 1)
. Administration of NBI-31772 increased Po by 34% at 10 days after injury (P < 0.05; Table 1
, Figure 1
). Administration of NBI-31772 also increased Pt by 26% and likewise increased sPo by 24% compared with untreated muscles 14 days after injury (P < 0.05; Table 1
). Administration of NBI-31772 hastened the time course of the twitch response throughout regeneration compared with untreated muscles, as evident from the 60, 23, and 26% increase in dPtwitch/dt at 10, 14, and 21 days, respectively (P < 0.05; Table 1
). Furthermore, NBI-31772 administration decreased time-to-peak twitch tension (TPT) by 15% and one-half relaxation time (
RT) by 29% compared with untreated TA muscles at 10 days after injury (P < 0.05; Table 1
). It is worth noting that administration of NBI-31772 enhanced functional recovery as evident from the higher Po values during regeneration (P < 0.05; main effect; Figure 1
).
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Representative H&E-stained transverse muscle sections and the median and individual CSA values for TA muscles 10, 14, and 21 days after myotoxic injury are presented in Figure 2
. The median CSA of myofibers was 35, 24, and 16% greater following NBI-31772 administration at 10, 14, and 21 days after injury, respectively (Figure 2)
.
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with embryonic MyHC isoform expression transitioning to MyHC IIa/x and MyHC IIb as regeneration progressed from 10 to 21 days (Figure 3)
. Administration of NBI-31772 increased the relative proportion of MyHC IIb with a concomitant decrease of MyHC IIa/x compared with untreated muscles at 21 days after injury (P < 0.05; Figure 3
).
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EDL and soleus muscle mass and diaphragm muscle strip thickness in mdx mice were not different following NBI-31772 treatment (Table 2)
. Administration of NBI-31772 did not alter the TPT responses in the soleus or diaphragm muscles of mdx mice but hastened RT in EDL muscles of mdx mice by 17% (P < 0.05; Table 2
). Administration of NBI-31772 induced a faster contractile phenotype in the EDL muscles as evident from the 26% increase in dPtetanic/dt in treated compared with untreated mdx mice (P < 0.05; Table 2
). NBI-31772 treatment also increased –dPtetanic/dt by 36% in diaphragm muscle strips compared with control mdx mice. In contrast to the data for regenerating TA muscles, NBI-31772 treatment did not alter Po or sPo of the EDL or soleus muscles or the sPo of diaphragm muscle strips.
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EDL muscles from NBI-31772-treated mdx mice were less susceptible to contraction-induced damage compared with muscles from untreated mdx mice (P < 0.05; Figure 4
). The force deficits following active stretches of 30, 40, and 50% beyond Lf were reduced by 18, 24, and 19%, respectively (P < 0.05; Figure 4
). Likewise, the diaphragm muscle strips from NBI-31772-treated mdx mice exhibited an overall protection from contraction-induced injury (P < 0.05, main effect; Figure 4
). The soleus muscles of mdx mice are already far less susceptible to contraction-induced injury than the EDL or diaphragm muscles and therefore demonstrated no change in injury susceptibility after NBI-31772 treatment (Figure 4)
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EDL muscles from NBI-31772-treated mdx mice were more fatigable and exhibited an impaired recovery of force producing capacity after repeated contractions such that following the fatigue protocol EDL muscles from NBI-31772-treated mdx mice produced 7% less force than control mdx mice (P < 0.05; Figure 5
). This NBI-31772-induced increase in EDL muscle fatigability was associated with a 14% decrease in citrate synthase activity (P < 0.05; Figure 5
). The soleus muscles from NBI-31772-treated mdx mice were also more fatigable and had impaired recovery of their force producing capacity compared with muscles from untreated mdx mice (P < 0.05; main effect, Figure 5
). Likewise, diaphragm muscle strips from NBI-31772-treated mdx mice produced 8% less force than untreated mdx mice after the fatigue protocol and also demonstrated an impaired functional recovery (P < 0.05; Figure 5
). Soleus muscles and diaphragm muscle strips did not exhibit any change in citrate synthase activity after NBI-31772 treatment (Figure 5)
.
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Representative H&E-stained transverse muscle sections and CSA data for the EDL, soleus, and diaphragm muscles are presented in Figure 6
. The median CSA of myofibers from EDL muscles of mdx mice was 12% smaller after NBI-31772 treatment compared with mdx mice (P < 0.05; Figure 6
). The median CSA of myofibers from soleus muscles of mdx mice was not different after treatment with NBI-31772 (Figure 6)
. Likewise, CSA of myofibers from the diaphragm muscles of mdx mice was not different after treatment with NBI-31772 (Figure 6)
. Administration of NBI-31772 did not alter the proportion of centrally nucleated fibers in the EDL, soleus, or diaphragm muscles from mdx mice (Table 2)
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Discussion |
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In regenerating TA muscles IGFBP inhibition increased force production and hastened the rates of contraction and relaxation. These functional improvements were associated (at day 21) with an increased proportion of MyHC type IIb. During muscle regeneration, there is a progression in MyHC isoform expression from early embryonic and neonatal isoforms through to mature MyHC isoforms such as type IIb and IIa/x.31 In uninjured TA muscles, 80% of the MyHCs are of the type IIb isoform, with the remaining 20% being MyHC type IIa/x.16 NBI-31772 treatment facilitated a more rapid progression from these slower-contracting MyHC isoforms to the faster MyHC isoforms and so increased the overall rate of regeneration. This is further evident from the increase in median fiber CSA observed in regenerating TA muscles after IGFBP inhibition.
Similar to its effects on regenerating TA muscles, IGFBP inhibition increased the speed of relaxation in EDL muscles from mdx mice, which was associated with a decrease in fatigue resistance, a reduced force producing capacity after fatigue, and a reduced citrate synthase activity. Taken together, these findings show that IGFBP inhibition caused a shift toward a faster, less oxidative muscle phenotype in mdx mice. This finding is in direct contrast to results obtained in previous studies where low-dose rhIGF-I administration resulted in a more oxidative, less fatigable muscle phenotype in mdx mice.14-16 However, following plasmid-based overexpression of IGF-I in rat skeletal muscle, a shift toward a more glycolytic phenotype has been observed,32 an effect also seen in transgenic mice over-expressing muscle-specific IGF-I.33 The effects of IGF-I on skeletal muscle oxidative capacity appear to be strongly modulated by IGFBPs, and IGFBP inhibition promotes a glycolytic muscle phenotype similar to that observed with IGF-I overexpression. These results also suggest that the skeletal muscle response to exogenous administration of rhIGF-I is altered through interactions with IGFBPs. Interestingly, in EDL muscles from dystrophic mdx mice, median fiber CSA decreased with IGFBP inhibition. Together with the effects on the regenerating TA muscle, this indicates that although IGFBP inhibition increases early events of muscle fiber regeneration, including proliferation and differentiation,8,9,34,35 it does not alter the rate of muscle fiber degeneration in mdx mice. Thus, for dystrophic muscles, IGFBP inhibition may increase the proportion of smaller caliber regenerating muscle fibers.
Similar to findings from our other studies where we administered rhIGF-I to mdx mice,16 IGFBP inhibition in mdx mice reduced the susceptibility to contraction-induced injury in EDL muscles and diaphragm muscle strips. This finding has significant therapeutic implications as contraction-mediated injury is a contributing factor to the progressive muscle fiber breakdown in muscular dystrophy3 and further highlights the therapeutic potential of modulating the IGF-I signaling pathway for ameliorating muscle pathologies. It is interesting to note that studies demonstrating a reduced susceptibility of dystrophic skeletal muscles to contraction-induced injury have also reported a shift toward a slower muscle phenotype. It has been shown that fast-twitch muscles from dystrophic mdx mice are particularly susceptible to contraction-induced damage,26,36,37 a finding supported by studies on Duchenne muscular dystrophy patients where fast-twitch fibers were more susceptible to such damage than slow-twitch fibers.38 From these studies, and from others by our laboratory,14-16 it had been hypothesized that the IGF-I-mediated protection from contraction-induced injury was due to the induction of a slower phenotype that renders muscles less susceptible to such injury. However, the results of the present study are the first to demonstrate protection from contraction-induced injury concomitant with a faster, less oxidative muscle phenotype. Although this increased fatigability may be viewed as being detrimental, it is unlikely that such repeated maximal contractions would ever be performed by patients. Regardless, in vivo measures of strength, endurance, and longevity would help to resolve these issues. Under normal situations, the primary disability in dystrophic muscles is an inability to generate sufficient force to perform the tasks of daily living because of a lack of viable muscle tissue, rather than a lack of functional capacity to perform sustained maximal contractions. It is therefore likely that the IGF-I-mediated protection from contraction-induced injury occurs through an altogether different mechanism, potentially by altering specific parameters of excitation-contraction coupling.16 From the contrasting results obtained with NBI-31772 and rhIGF-I administration in mdx mice, it is clear that IGFBPs play a significant role in the determination of skeletal muscle phenotype, either independently or via their interactions with IGF-I.
We have demonstrated that systemic inhibition of IGFBPs has beneficial effects for muscle fiber regeneration and for the dystrophic pathology in mdx mice. One of the limitations of the present study was the use of a non-specific aptamer that inhibited the actions of all six IGFBPs. Previous studies have demonstrated that during regeneration skeletal muscle levels of IGFBPs are regulated differentially. For example, IGFBP3 expression is increased during early muscle regeneration following ischemic injury,39 whereas IGFBPs 4, 5, and 6 are increased during the later stages of regeneration.39,40 Future studies should aim to inhibit specific IGFBPs to determine the exact role of each isoform in skeletal muscle regeneration. Identifying the specific roles for IGFBPs during muscle fiber regeneration will have important therapeutic implications for the muscular dystrophies and muscle wasting disorders.
In conclusion, this study examined the effect of modulating IGFBP interactions with IGF-I on the functional properties of regenerating skeletal muscle and on the dystrophic pathology in mdx mice. We have demonstrated an entirely novel way of manipulating IGF-I signaling for improving muscle fiber regeneration. We demonstrated that IGFBP inhibition significantly enhanced muscle fiber regeneration and increased the rate of functional recovery after myotoxic injury. We also showed that IGFBP inhibition in mdx mice reduced the susceptibility of dystrophic muscles to contraction-mediated injury, a contributing factor to the etiology of the dystrophic pathology. The findings highlight the therapeutic potential of IGF-I signaling for pathologies where muscle wasting and weakness are indicated.
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Footnotes |
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Supported by research grants from the Muscular Dystrophy Association (USA).
J.D.S. and S.M.G. contributed equally to this work.
Accepted for publication June 19, 2007.
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References |
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