Merosin-deficient congenital muscular dystrophy type 1A (MDC1A) is a severe and fatal muscle-wasting disease with no cure. MDC1A patients and the dyW−/− mouse model exhibit severe muscle weakness, demyelinating neuropathy, failed muscle regeneration and premature death. We have recently shown that laminin-111, a form of laminin found in embryonic skeletal muscle, can substitute for the loss of laminin-211/221 and prevent muscle disease progression in the dyW−/− mouse model. What is unclear from these studies is whether laminin-111 can restore failed regeneration to laminin-α2-deficient muscle. To investigate the potential of laminin-111 protein therapy to improve muscle regeneration, laminin-111 or phosphate-buffered saline-treated laminin-α2-deficient muscle was damaged with cardiotoxin and muscle regeneration quantified. Our results show laminin-111 treatment promoted an increase in myofiber size and number, and an increased expression of α7β1 integrin, Pax7, myogenin and embryonic myosin heavy chain, indicating a restoration of the muscle regenerative program. Together, our results show laminin-111 restores muscle regeneration to laminin-α2-deficient muscle and further supports laminin-111 protein as a therapy for the treatment of MDC1A.
Merosin-deficient congenital muscular dystrophy type 1A (MDC1A) is a devastating genetic disease that results in muscle weakness from birth. MDC1A is caused by mutations in the LAMA2 gene, which results in loss of laminin-211/221 (merosin) from the basal lamina of skeletal and cardiac muscle (1). MDC1A patients experience muscle atrophy, impaired muscle regeneration, increased muscle apoptosis, fibrosis and progressive muscle loss (2,3). Most MDC1A patients are severely affected within the first year of life and never achieve independent ambulation (2). As the disease progresses, MDC1A patients exhibit joint contractures, respiratory complications, scoliosis, feeding difficulties, limited eye movement, dysmyelinating neuropathy and seizures (4–7). Palliative interventions such as physical therapy, cough assist and spinal fusion are the only available treatments, yet MDC1A patients remain susceptible to respiratory failure and premature death as early as the first decade of life (5,8).
Understanding of the pathophysiology of MDC1A has led to novel treatment approaches including anti-apoptotic therapies, regeneration enhancers and restoration of the missing basal lamina. Studies using mouse models of MDC1A have shown that transgenic expression of laminin-α1 (9–11), laminin-α2 (12,13), mini-agrin (14,15), GalNAc transferase (16), insulin-like growth factor 1 (17), α7 integrin (18) and Bcl-2 (19) can reduce or prevent disease progression. Pharmacological interventions with doxycycline and omigapil, which inhibit apoptotic pathways, and 3-methyladenine, which blocks autophagy, have also been shown to improve preclinical outcomes including improved survival and reduced muscle pathology (3,19–21). Together, these studies indicate therapies that restore interactions between the muscle and extracellular matrix and/or normalized muscle survival signaling pathways may be beneficial in the treatment of this devastating muscle-wasting disease.
The regenerative capacity of muscle is dependent on the activation and proliferation of normally quiescent satellite cells followed by their subsequent differentiation into mature myofibers. Satellite cells are located proximal to muscle fibers within the laminin-rich basal lamina and become activated by cues induced by muscle injury or disease. Signaling between laminins and their cognate receptors such as the α7β1 integrin are among the environmental cues necessary for satellite cells to proliferate and repair damaged muscle. Loss of laminin-211/221 in MDC1A patients and MDC1A mouse models also results in a secondary loss of the α7β1 integrin and a reduced ability of satellite cells to be activated, proliferate and support efficient muscle repair (13,22). Together, these observations indicate the importance of the laminin-rich microenvironment for muscle repair.
Recently, we have shown that laminin-111 can act as a protein substitution therapy for laminin-α2 deficiency in mice (23). Our studies showed that dyW−/− mice treated with laminin-111 exhibit reduced muscle pathology, apoptosis, fibrosis, maintained muscle strength and mobility and dramatically increased lifespan (23). What is unclear from these studies is whether the therapeutic effect of laminin-111 is due, in part, to a restoration of the muscle regeneration program.
In this study, we examined muscle repair in laminin-111-treated dyW−/− muscle following cardiotoxin (CTX)-induced muscle injury. Our results show laminin-111 treatment improved the timing and extent of muscle repair and regeneration in laminin-α2-deficient muscle, and provides further evidence for the therapeutic potential of laminin-111 protein therapy for MDC1A and other muscle diseases in which regeneration is defective.
Laminin-111 improves muscle repair in laminin-α2-deficient mice after CTX-induced injury
Recent studies have shown that laminin-111 protein therapy can substitute for the loss of laminin-211/221 and ameliorate the progression of disease in the dyW−/− mouse (23). One pathological component of laminin-α2 deficiency is a profound delay in the ability to regenerate skeletal muscle (14,15). The muscle regeneration that occurs in various muscular dystrophies can be ongoing and to quantify any deficiencies in regeneration, toxins such as CTX or notexin are often used to reset the cycle of degeneration and regeneration to a defined moment. To investigate whether the therapeutic effect of laminin-111 is also due to restored muscle regeneration, we pretreated TA muscle of laminin-α2 null mice with either laminin-111 or PBS, artificially induced muscle degeneration with CTX three days later, and assessed the timing and extent of muscle regeneration using histological and morphological measurements.
Three-week-old dyW−/− mice (Day 3) were injected intramuscularly (i.m.) with 100 µL of 1500 nM Engelbreth-Holm-Swarm (EHS) laminin-111 or PBS. Three days later, the tissue from Day 0 mice was harvested (Fig. 1A), and mice for subsequent Day 4 and 10 analyses were injected i.m. with CTX to induce muscle damage (Fig. 1A). To ensure delivery of EHS laminin-111 to TA muscles, cryosections were stained for laminin-α1 and imaged at 400× using high-resolution confocal microscopy (Fig. 2A–F). Following laminin-111 treatment, the laminin-α1 chain was localized within the basal lamina of all TA muscles injected with laminin-111. No laminin-α1 was evident in PBS-treated muscles, consistent with previous reports (23).
During muscle regeneration, increased synthesis of the contractile apparatus within nascent myotubes increases the myofiber cross-sectional area (CSA, Fig. 2G–L). While CSA of myofibers is dependent on fiber orientation and should be used cautiously, the high number of myofibers quantitated and the small error bars indicate that in this case the measurements are valid. Muscle cryosections stained by hematoxylin and eosin (H&E) demonstrated that by Day 0 (before CTX injection) laminin-111-treated TA muscle demonstrated an average myofiber CSA of 1032 µm2, whereas myofibers treated with PBS were 988 µm2 (Fig. 2G, J and M). By Day 4 (4 days after CTX injection), laminin-111- and PBS-treated muscle demonstrated an average CSA of 1803 and 1442 µm2, respectively; Fig. 2H and 2K. At 10 days after CTX injection, the average CSA was lower in both treatment groups, 877 µm2 for laminin-111 and 769 µm2 for PBS (Fig. 2I, L and M).
Compared with PBS-treated muscle, the increase in average CSA was marginally higher in laminin-111-treated muscle at Day 0 and had increased 25% by Day 4. The average CSA in laminin-111-treated muscle had declined 10 days after CTX to a 14% increase in CSA compared with PBS-treated muscle (Fig. 2M).
The lack of sustained effect in average and peak CSA in laminin-111-treated muscle, following CTX injury, may be attributable to the severity of disease in this mouse model or the need for multiple laminin-111 treatments. The artificially harsh conditions and inflammation following CTX injury may have also accelerated the degradation of injected laminin-111.
Not only was the average and peak size of myofibers increased after laminin-111 therapy and CTX injury, there was also a dramatic 100% increase in the number of myofibers by Day 4 (Fig. 2N). By Day 10, when comparable values for average and peak CSA were noted in laminin-111 and PBS-treated muscles, laminin-111 treatment resulted in an 800% increase in the number of myofibers (Fig. 2N). These results indicate that the therapeutic effect of laminin-111 protein substitution in laminin-α2-deficient muscle is due, in part, to improved size of individual myofibers, but also increased survival of laminin-α2-deficient muscle.
Laminin-111 increases muscle regeneration in laminin-α2-deficient muscle
Embryonic myosin heavy chain (eMyHC) is transiently expressed during muscle development and is used as a marker for nascent myofiber creation during muscle regeneration. As myofibers further differentiate, eMyHC becomes replaced by adult myosin heavy chain (23). On Day 0, before CTX injury, laminin-α2-deficient TA muscle injected with PBS demonstrated that 14% of myofibers were eMyHC positive (Fig. 3A), indicating that persistent regeneration is ongoing in dyW−/− muscle. By Day 4, following CTX injection, 54% of PBS-treated myofibers were eMyHC positive, which had decreased to 9% by Day 10 – levels that were observed pre-CTX (Fig. 3B and C).
In contrast, muscles treated with laminin-111 at Day 0 (before CTX injury) demonstrated 31% eMyHC-positive myofibers, indicating that laminin-111 treatment rapidly increases the pool of differentiating myofibers (Fig. 3D). By Day 4, the number of eMyHC-positive myofibers in laminin-treated muscle had increased to 86% of all myofibers, representing a 59% increase above PBS-treated muscles (Fig. 3E and J). By Day 10, eMyHC-positive myofibers in laminin-111-treated muscle had declined to 16%, representing a 77% increase above PBS treatment levels (Fig. 3F and J). The biphasic appearance of eMyHC myofibers in both PBS- and laminin-111-treated muscles is consistent with the transient expression of eMyHC in regenerating myofibers and that myofiber differentiation was near completion. The increase in eMyHC-positive myofibers following laminin-111 treatment (even before CTX injury) demonstrates that laminin-111 protein increases the regenerative capacity of laminin-α2 null muscle.
We next examined whether the improved muscle regeneration as a result of laminin-111 treatment resulted in larger eMyHC-positive myofibers. To examine this, we measured the area of eMyHC-positive myofibers in wild-type TA muscle treated with PBS and dyW−/− TA muscles treated with either PBS or 1500 nM laminin-111. At Day 0, the average CSA of eMyHC fibers in dyW−/− muscle following treatment with PBS and laminin-111 was 279 and 360 µm2, respectively (Fig. 4A). Wild-type TA muscle possess a normal repair capacity, and the average CSA of eMyHC-positive myofibers of 910.0 µm2 following PBS treatment was much larger than dyW−/− muscle treated with PBS or laminin-111 (Fig. 4A). At Day 0, there was no statistical difference in the peak CSA of eMyHC fibers either treated with PBS or laminin-111 (Fig. 4B), and both were only 42% the size of wild-type fibers prior to injury.
At 4 days after CTX, the average and peak CSA of myofibers from dyW−/− mice treated with laminin-111 approached wild-type levels (Fig. 4C and D). The average CSA of eMyHC-positive fibers treated with laminin-111 were 89% of the average CSA of wild-type PBS-treated eMyHC fibers (Fig. 4C). Peak CSA of eMyHC fiber treated with laminin-111 was indistinguishable from the CSA observed in wild-type mice (Fig. 4D). The average CSA for PBS-treated tissue, laminin-111-treated tissue and wild-type tissue was 283, 363 and 410 µm2, respectively (Fig. 4C). Laminin-111-treated laminin-α2 null muscle had a peak CSA of 251 µm2, whereas PBS-treated tissue was 153 µm2 (Fig. 4D). The area of eMyHC fibers at peak CSA treated with PBS was 60.7% the area of peak eMyHC found in wild-type muscle, whereas tissue treated with laminin-111 was nearly identical (250.4 µm2 in laminin-111-treated versus 252.2 µm2 in PBS-treated, Fig. 4D). Compared with PBS-treated dyW−/− muscle, the improvement in average CSA of eMyHC-positive myofibers following laminin-111 treatment seen at Day 4 was only minimally evident by Day 10, and there was no statistical difference of peak CSA of eMyHC fibers (Fig. 4E and F). The decline in the efficacy of laminin-111 seen at Day 10 may be due to rapid matrix turnover in muscle. Nonetheless, the data shows that laminin-111 treatment restored regenerative capacity to laminin-α2-deficient muscle prior to CTX-induced injury by increasing the percentage and the average CSA of eMyHC fibers.
Laminin-111 increases α7β1 integrin in CTX-damaged dyW−/− muscle
The α7β1 integrin is a laminin receptor and is expressed on satellite cells and adult muscle (24–26). The α7 integrin knockout mouse exhibits reduced laminin-α2 expression and upon CTX challenge has a defect in muscle regeneration that is similar to the dyW−/− and dy3k mouse models of MDC1A (24,25). Loss of laminin-α2 in dyW−/− and dy3K mouse models also results in a secondary reduction of sarcolemmal localization of α7β1 integrin, which likely exacerbates the muscular dystrophy phenotype (22,27,28). Therefore, laminins and the α7β1 integrin have a cooperative relationship that is critical for effective muscle regeneration and sarcolemmal integrity (18,23,29,30).
To examine whether laminin-111 protein therapy could restore sarcolemmal localization of the α7β1 integrin in laminin-α2 null muscle, integrin localization was examined in TA muscles treated with 1500 nM laminin-111 or PBS before and after CTX damage. Immunofluorescence revealed TA muscle treated with laminin-111 before or after CTX had an increase in sarcolemmal localization of both α7 and β1D integrin compared with PBS-treated muscle (Fig. 5A–I). The results indicate treatment with laminin-111, a high affinity ligand for the α7β1 integrin, increased the membrane localization of the α7β1 integrin in laminin-α2-deficient muscle. These data are consistent with earlier data in which transgenic laminin-α1 expression in dy3k mice restores sarcolemmal α7β1 integrin (31). In addition, increased levels of α7β1 integrin in TAs treated with laminin-111 were more uniform in size and had less atrophic muscle fibers.
To confirm our immunofluorescence results, western analysis of the α7A, α7B and β1D integrins was performed on TA tissue treated with PBS or laminin-111 either prior to CTX damage (Day 0) or 4 and 10 days after CTX-induced damage. From Days 0, 4 and 10 day time points, laminin-111 treatment resulted in greater protein levels of α7A, α7B and β1D: 1.8-, 1.4- and 2.0-fold increase in α7A, α7B and β1D integrin protein, respectively, at Day 0, a 1.7-, 1.2- and 2.1-fold increase in α7A, α7B and β1D integrin, respectively, at Day 4 and a 1.5-, 1.3- and 1.8-fold increase in α7A, α7B, β1D integrin, respectively, at Day 10 (Fig. 5J–L). Together, this data show that laminin-111 treatment in dyW−/− muscle before damage stabilizes the α7β1 integrin at the sarcolemma. Given the presence of α7B on satellite cells and myofibers, and the presence of α7A and β1D on mature fibers only, the laminin-111 treatment appears to target both satellite cells and mature myofibers.
Laminin-111 treatment increased total TA area in laminin-α2-deficient muscle
We previously demonstrated that as few as three weekly systemic intraperitoneal injections of laminin-111 to dyW−/− mice resulted in more individual myofibers in the triceps brachii and a larger overall muscle area, indicating that laminin-111 promotes de novo muscle generation and/or is mitigating the loss of myofibers owing to laminin-α2 deficiency. To clarify this result, we quantified the total average TA area at all timepoints and treatments. At Days 0, 4 and 10, the average total area of TA muscles treated with PBS was 65 000, 57 500 and 73 600 µm2, respectively whereas the average total area of TA muscles treated with laminin-111 was 107 000, 70 000 and 117 000 µm2, respectively (Fig. 6G). Overall, all TAs treated with laminin-111, before or after CTX damage, were larger than PBS controls resulting in a 65, 22 and 59% increase in total TA area at Days 0, 4 and 10, respectively (Fig. 6G, Supplementary Material, Fig. S1). The results substantiate our earlier findings and suggest that laminin-111 promotes de novo muscle generation in laminin-α2 deficiency.
Laminin-111 treatment reduces fibrosis in laminin-α2-deficient muscle
The persistent cycles of degeneration and regeneration that occur in muscular dystrophies, including MDC1A, are also accompanied by chronic inflammation, which ultimately results in the deposition of fibrotic matrices (18,23,32). In order to determine the effects of laminin-111 treatment on fibrosis in laminin-α2-deficient muscle before and after CTX-induced injury, TA muscle was stained with Sirius Red as previously described (33,34). The fibrotic area of PBS-treated TA muscle at Days 0, 4 and 10 was 15, 56 and 22%, respectively (Fig. 6A–C and H, Supplementary Material, Fig. S1). In contrast, treatment with laminin-111 reduced these fibrotic areas in the TA muscle at Days 0 and 4 to 8.5 and 39%, respectively (Fig. 6D, E and H). However, by Day 10, the fibrotic area of PBS- and laminin-111-treated TA muscle was similar (Fig. 6C, F, H).
These results indicate that laminin-111 has an immediate impact on muscle fibrosis and following CTX injury can mitigate further fibrotic accumulations in laminin-α2-deficient muscle. However, by 10 days, the anti-fibrotic effect of laminin-111 had waned.
Laminin-111 increases Pax7 and myogenin in laminin-α2 null muscle after damage
Satellite cells express the paired box transcription factor Pax7 and are normally quiescent. Induced to proliferate by muscle damage or disease, satellite cells continue to express Pax7 in addition to the myogenic regulator factor (MRF) MyoD. As satellite cells further differentiate into myotubes, Pax7 expression is reduced and MyoD expression is replaced by expression of the MRF myogenin (35–37). Thus, Pax7 and myogenin mark the early and late stages of satellite cell-based muscle regeneration, respectively, and were measured to ascertain whether laminin-111 enhanced the early or late stages of myogenesis in laminin-α2-deficient muscle.
Compared with PBS-treated muscle, laminin-111-treated muscle resulted in a 1.7-, 2.4- and 1.4-fold increase in Pax7 protein 0 (before CTX), 4 and 10 days after CTX injection (Fig. 7A). The decline in Pax7 at Day 10 in PBS- and laminin-111-treated groups is consistent with terminal differentiation. At Day 0, prior to CTX injury, there were no significant differences in myogenin expression in laminin-111 or PBS-treated tissue (Fig. 7B), indicating that laminin-111 is acting upon recently activated (myogenin negative) satellite cells rather than nascent myotubes exhibiting aborted terminal differentiation (24). Following CTX, however, laminin-111-treated muscle exhibited a 2.5- and 3-fold increase in myogenin at Day 4 and 10, respectively, compared with PBS treatment (Fig. 7B), indicating that laminin-111 increased the pool of satellite cells available for terminal differentiation. The increase in Pax7 and myogenin following laminin-111 treatment and CTX injection is consistent with the positive mitogenic effect laminin-111 is known to provide to proliferating myoblasts (24).
Mobility is improved in dyW−/− mice in which the TA muscles were treated with laminin-111
Mobility observations of dyW−/− mice treated once with either laminin-111 or PBS i.m. injected and injured 3 days later with CTX demonstrated that animals treated with laminin-111 exhibited more effective use of the treated leg than the leg treated with PBS (Supplementary Material, Fig. 2S). It was also observed that the leg of mice receiving i.m. treatments with laminin-111 in the TA muscle showed decreased joint contractures and hind limb neuropathy than PBS-treated dyW−/− mice of the same age (Supplementary Material, Fig. S2). Together, these data show that laminin-111 improved muscle repair of the TA muscle, despite damage by CTX, and this translated into improved overall muscle function after CTX-induced damage.
Weekly laminin-111 therapy sustains muscle regeneration in laminin-α2-deficient muscle
The effectiveness of a single dose of laminin-111 on regeneration of laminin-α2-deficient muscle had waned 10 days after CTX injection (Fig. 2H). Activation of matrix metalloproteases in laminin-α2-deficient muscle may have increased the turnover of active laminin-111 in muscle by Day 10, and in this study, inflammation as a result of the CTX injection may also have promoted further turnover of the injected laminin-111. To test this idea, additional laminin-111 protein was injected at 4, 11, 18 and 25 days after CTX damage in order to boost the amount of active laminin-111 in the muscle (Fig. 1B, B = Booster treatments with laminin-111).
There was only a 44 µm2 difference in average eMyHC CSA between treatment groups at Day 10 with one dose of laminin-111 (Fig. 4E). Day 10B TAs that received a boost of laminin-111 at 4 days after CTX (Fig. 8C: Day 10B) resulted in a 360 µm2 increase in average CSA, representing an 8.3-fold increase versus treatment groups treated only once prior to injury (Fig. 8A). The increase in average CSA observed at Day 10 following weekly injections continued to be evident at Day 28 (Fig. 8A).
By 10 days after CTX injury, the peak eMyHC CSA of muscle treated once with laminin-111 was nearly identical to the PBS-treated fibers. However, by 10 days after CTX injury, those muscles injected weekly with laminin-111 exhibited a peak CSA area which was significantly higher than the PBS-treated tissue (Figs 4D, 8D). Furthermore, by Day 28 after CTX, the peak CSA area in both PBS and laminin treated muscles exhibited a decrease yet the peak CSA area in laminin treated muscle remained larger than PBS treated muscle (Fig. 8B). In addition to having larger fibers in the laminin-111-treated muscles, there was also a greater number of fibers compared with the PBS-treated tissue (Fig. 9C).
There was a similar trend in total TA area observed in both Day 10 treatment groups (Fig. 6G and Fig. 9A). The total TA area of Day 10B weekly laminin-111-treated tissue was 1.3-fold larger than those treated with PBS and 1.5-fold larger at Day 28B. Total TA area at Day 28 with weekly treatments of laminin continued to be larger than controls and was also larger than either of single treatment Day 10 laminin-treated TAs (Figs 9A, 6G, Supplementary Material, Fig. S1). The improvements with weekly laminin-111 injections are also evident in further reduced fibrosis at 10 and 28 days after CTX injury. Although no statistical difference was measured in percent fibrosis in the Day 10 mice that received only one treatment, there was a significant difference in both the Day 10 weekly injected group and the Day 28 group (Figs 9B, 6H). The percent fibrosis of the PBS weekly injected mice increased from Day 10B to Day 28B. This result is expected owing to the severe progressive nature of this disease; however, it was surprising to see a continued decrease in fibrosis in the laminin-111-treated tissue from Day 10B to Day 28B (Fig. 9B). These data provide evidence that weekly injections in laminin-α2-deficient muscle are necessary to maintain the efficacy of laminin-111 in muscle regeneration and indicate the half-life for laminin-111 activity in laminin-α2-deficient muscle under these experimental conditions of between 3–7 days.
MDC1A is a progressive muscle-wasting disease caused by the absence of laminin-211/221 owing to defects in the LAMA2 gene. Laminin-211/221 is a component of the basal lamina and critical for normal muscle function. There is no cure or approved therapy for MDC1A and only palliative treatment options are available. Recently, we have shown that systemic delivery of laminin-111 protein targets skeletal muscle and reduces the progression of disease in the dyW−/− mouse model of MDC1A (23).
In the current study, we show that laminin-α2-deficient muscle treated with laminin-111 protein before CTX injury increased the regenerative capacity of skeletal muscle; measured as increased average CSA, α7β1 integrin, Pax7 and myogenin expression and ultimately increased myofiber number. Importantly, laminin-111 appears to increase de novo myogenesis.
This result is consistent with previous in vitro and in vivo studies, which show increased contact between satellite cells, their proximal myofibers and the extracellular matrix increases satellite cell proliferation and migration and decreases programmed cell death, myofiber loss, fat deposition and fibrosis in the laminin-α2-deficient muscle (13,14,38). The peak activity of PBS and laminin-111 protein treatment for myogenic repair and regeneration in laminin-α2-deficient muscle was 4 days after CTX damage. At 10 days after CTX damage, the repair capacity in laminin-α2 null muscle was diminished, indicating reduced myogenic activity of laminin-111 in the muscle or alternatively a completed cycle of differentiation.
The reduced activity of a single injection of laminin-111 10 days after CTX injection is likely due to the accelerated turnover of the extracellular matrix by proteases and inflammatory infiltrates during degeneration. The α7β1 integrin and α-dystroglycan are the predominant laminin receptors on adult skeletal muscle. This study showed that laminin-111 protein therapy is able to restore sarcolemmal integrity in laminin-α2-deficient muscle though increased levels and sarcolemmal localization of α7β1 integrin. Sarcolemmal α-dystroglycan is not appreciably reduced in dyW−/− animals, yet the interaction of laminin-111 with the dystroglycan–dystrophin glycoprotein complex likely synergizes with the α7β1 integrin to reinforce muscle integrity (29,30). The affinity of α7β1 integrin for laminin-111 has been shown to be stronger than α-dystroglycan (39–42). The increased level of α7β1 integrin affects not only sarcolemmal organization but also the earlier stages of myogenesis such as satellite cell proliferation and migration; and ultimately the regenerative capacity of skeletal muscle (25,35).
Mini-agrin also promotes regeneration in MDC1A mouse models but appears to do so without enhancement of the α7β1 integrin (15). Importantly, the improvement in muscle regeneration in the dyW−/− mouse by mini-agrin is not accompanied by an increase in the number of myofibers as seen following laminin-111 treatment (43). The α7 integrin KO mouse exhibits a mild myopathy but is otherwise healthy and fertile (24). However, following CTX injection into the TA, the α7 integrin KO mouse exhibits a profound defect in muscle regeneration that mimics what occurs in MDC1A mouse models (24). Despite the absence of α7 integrin and a mildly reduced composition of laminin-211/211, the injected laminin-111 was also able to rescue the regenerative defect in α7 KO mice. These observations indicate that existing α-dystroglycan perhaps with assistance from up-regulated α6β1 integrin can rescue the regenerative defect in these mice. Overall, our results indicate that matrix interactions are generally important for muscle regeneration, but the cooperative interaction between α-dystroglycan and the α7β1 integrin, perhaps at differing stages of myogenesis, is crucial for normal muscle regeneration.
Previous research has shown that changes in α7β1 integrin expression in dyW−/− muscle can alter the organization/deposition of extracellular matrix proteins and enzymes that regulate those proteins (18). These results indicate that increased α7β1 integrin owing to laminin-111 protein therapy may be a major contributing factor to sarcolemmal and basal lamina stability.
The increased number of eMyHC-positive myofibers in laminin-α2 null muscle after laminin-111 treatment observed in this study demonstrates the ability of laminin-111 protein therapy to improve the repair capacity of existing muscle and provide an environment for de novo muscle formation (14,33,35). Our results indicate laminin-111 protein therapy not only decreases the pathology of laminin-α2-deficient mice (23) but also improves the timing, rate and repair capacity, which may explain improvements in muscle disease observed with systemic delivery of the protein.
This supports the idea that laminin-111 may confer a protective effect by providing new mechanical linkages between the extracellular matrix and the sarcolemma (23). This is consistent with other studies, which show that loss of contact between myofibers through the basal lamina and extracellular matrix initiates programed cell death in laminin-α2-deficient muscle and that the restoration of contact through laminin-111 treatment in dyW−/− mice leads to reduction of apoptosis, muscle degeneration and myofiber loss (14).
The myogenesis and regenerative capacity of skeletal muscle is dependent on the interactions of satellite cells in muscle and the laminin-rich basal lamina (33,35,44). During activation, satellite cells express transcription factors Pax3, Pax7, MyoD, myogenin and MRF4. We have previously shown that the loss of α7 integrin leads to reduced satellite cell activation and myoblast differentiation in response to muscle injury (18,23). In this study, we demonstrate laminin-111 treatment results in increased expression of Pax7 and myogenin, which are markers for satellite cell activation and terminal myogenic differentiation, respectively, (26,37,45). In muscle receiving one treatment with laminin-111, Pax7 and myogenin were dramatically increased at Days 4 and 10, indicating an increase in satellite cell activation and myogenic repair capacity. These results indicate laminin-111 can act to replace the loss of laminin-211/221 in the basal lamina to improve satellite cell activation and repair.
The effect of laminin-111 treatment was also visually apparent by observing the mobility of dyW−/− mice treated with laminin-111 or PBS. The movement and use of the legs in which the TA muscle was treated with laminin-111 and then subjected to CTX damage was closer to wild-type, whereas the legs in which the TA muscle was treated only with PBS and subjected to damage exhibited reduced mobility and use. This result is consistent with our previous study demonstrating improved mobility and activity following systemic treatment with laminin-111.
Together, this study indicates that laminin-111 protein therapy possesses two mechanisms of action that are relevant to normal muscle function but also necessary for effective treatment of MDC1A;  restoration of the defective basal lamina, reinforced muscle adhesion of existing muscle and mitigation of the secondary manifestations of laminin-α2 deficiency, and  restoration of the laminin–integrin microenvironment is necessary for the expansion and terminal differentiation of satellite cells and thus effective muscle regeneration. Given the importance of the laminin–integrin relationship in muscle regeneration and adult muscle stability, laminin-111 protein therapy may also benefit other muscle diseases that exhibit defective muscle repair including Fukuyama muscular dystrophy, Fukutin-Related Muscular Dystrophy (LGMD2I), dysferlinopathy (LGMD2B) and dystroglycanopathy (MDC1D). In addition, laminin-111 protein therapy may be useful in the treatment of severe muscle injury and trauma or muscle loss associated with aging or chronic disease.
MATERIALS AND METHODS
All experiments involving mice were performed under an approved protocol from the University of Nevada, Reno Institutional Animal Care and Use Committee. The dyW+/− mice were a gift from Eva Engvall via Paul Martin (The Ohio State University, Columbus, OH, USA). For this study, dyW+/− mice, which are heterozygous at the lama2 locus, were breed to produce male dyW−/− and dyW+/+ (wild-type) animals, which were then used experimentally. Experimental procedures were performed once mice were 21 days of age. To reduce experimental bias, investigators assessing and quantifying experimental outcomes were blinded to the treatment and control groups following recently published guidelines (46).
Engelbreth-Holm-Swarm-derived natural mouse laminin-111 (Invitrogen Life Technologies, Grand Island, NY, USA) was thawed overnight at 4°C. At Day 3, mice were injected intramuscularly (i.m.) into the left tibialis anterior (TA) muscle with either 100 µL sterile PBS or 100 µL 1500 nM EHS laminin-111 in PBS. The right TA muscles were injected with 100 µL sterile phosphate-buffered saline (PBS) and served as controls. Day 0 mice were not injected with CTX and served as non-injury controls (Fig. 1A and B). The muscles were harvested either at 0, 4, 10 or 28 days after CTX injection for analysis.
CTX-induced muscle injury
TA muscles were damaged at Day 0, three days after laminin or PBS treatments by i.m. injection of 100 µL of a 10 µmol/L CTX solution (C3987; Sigma, St. Louis, MO, USA) in PBS. At 4, 10, or 28 days after CTX-induced injury, the mice were euthanized and TA muscles harvested for analysis. Skeletal muscles were dissected and flash-frozen in liquid nitrogen-cooled isopentane. The tissues were stored at −80°C until used for analysis.
Hematoxylin and eosin (H&E) staining of TA muscle was done as previously described (24). H&E-stained slides were used for Day 0, 4 and 10 analyses. Average CSA and total number of myofibers were counted at 200× magnification under bright-field microscopy using a Zeiss Axioskop 2 Plus fluorescent microscope, Zeiss AxioCam HRc digital camera and Axiovision 4.8 software. The total number of muscle fibers was determined by counting fibers in a minimum of six fields of view. A minimum of 800 muscle fibers per TA muscle were counted, with at least 4 TA muscle sections from each genotype, treatment and time point represented. Analysis of average CSA was quantified from a minimum of 3100 muscle fibers per group per time point and was quantified as previously described (24). Results are reported as the average fiber CSA of all muscle fibers circled for each time point and treatment. CSA was measured using Axiovision 4.8 software. All the areas in the peak increment were used to determine the average peak fiber area.
TA muscle sections were stained with Sirius Red to measure fibrosis in the muscle tissue. Sections on slides were fixed in 100% ethanol and then hydrated through an alcohol series (95 and 80% ethanol) and rinsed in tap water. The sections were stained with Sirius Red (0.1% in saturated aqueous picric acid solution, Rowley Biochemical Institute, Danvers, MA, USA) for 30 min followed by two washes in acidified water. The sections were dehydrated through an alcohol series, rinsed in xylene and mounted with DEPEX Mounting media (Electron Microscopy Science, Hatfield, PA, USA) (47). Images of each TA for each time point and treatment group were captured and analyzed using Axiovision 4.8 software. Areas of red in the TA were considered fibrotic. Circled fibrotic areas were added together, and any non-fibrotic fibers within the fibrotic area were subtracted from the calculated area. The percentage of muscle fibrosis was quantified in treated and control muscles as a percentage of total TA muscle area.
Sirius red slides were used for average and peak CSA calculations of Day 10B and Day 28B (B = Booster treatments with laminin-111) from mice injected weekly with laminin-111. Three composite images representing three whole TA sections taken at 100× magnification were used for analysis. All muscle fibers within an entire TA were used to determine the average CSA and total number of myofibers.
TA muscles were embedded in Tissue-Tek OCT, and 10-µm cryosections were cut using a Leica CM 1850 cryostat (Leica, Wetzlar, Germany). The sections were placed on pre-cleaned Surgipath slides (Surgipath Medical Industries, Richmond, IL, USA) and fixed using methanol, acetone and/or 4% paraformaldehyde. The Mouse on Mouse (M.O.M.) kit was used with all mouse antibodies according to package instructions (FMK-2201, Vector Laboratories Inc., Burlingame, CA, USA). Laminin-α1 chain was detected using a rat anti-mouse laminin-α1 monoclonal antibody (MAB1903; EMD Millipore Corporation, Billerica, MA, USA; 1:50) overnight followed by a fluorescein isothiocyanate (FITC)-conjugated goat anti-rat-IgG secondary antibody (1:5000; Li-Cor Biosciences, Lincoln, NE, USA).
Embryonic myosin heavy chain (eMyHC) was detected using bovine anti-mouse myosin heavy chain 2B antibody (BF-F3; Developmental Studies Hybridoma Bank, Iowa City, IA, USA; 1:30) overnight followed by FITC-conjugated anti-mouse-IgG secondary antibody. These slides were also treated with tetramethylrhodamine-labeled wheat germ agglutinin (WGA, 1:250, Molecular Probes, Invitrogen detection technologies, Eugene, OR, USA). The percentage of eMyHC-positive myofibers, average CSA and peak CSA eMyHC was calculated for each time point and treatment group.
The α7 integrin was detected using the rat monoclonal antibody CA5.5 (1:1000; Sierra Biosource, Morgan Hill, CA, USA) for 1 hour at room temperature. β1D integrin was detected using the mouse anti-mouse β1D integrin monoclonal antibody (1:25; MAB1900; EMD Millipore Corporation) overnight followed by a FITC-conjugated anti-mouse-IgG secondary antibody (1:5000; Li-Cor Biosciences). Slides were mounted using Vectashield Hard Set with DAPI (Vector Laboratories Inc.).
Images were captured using a Zeiss Axioskop 2 Plus fluorescent microscope, Zeiss AxioCam HRc digital camera and Axiovision 4.8 software or an Olympus FluoviewFV1000 Laser scanning biological confocal microscope using the Olympus micro FV10-ASW 3.1 software. Representative images for publication were taken at 400× using the Olympus FluoviewFV1000 Laser scanning biological confocal microscope.
The TA muscles from male mice were dissected, macerated and protein-extracted in RIPA as previously described (23). Protein was quantified using the Pierce BCA Protein Assay kit (Thermo Scientific, Rockford, IL, USA) according to manufacturer's directions and separated using 8 or 16% SDS PAGE and transferred to nitrocellulose membranes. The α7B integrin was detected with a 1:1000 dilution of rabbit anti-α7B (B2 347) polyclonal antibody overnight. α7A integrin was detected using a 1:1000 dilution of rabbit anti-α7A (CDB 345) antibody overnight. The β1D integrin was visualized using a 1:1000 rabbit anti-β1D-antibody (a gift from Woo Keun Song, Gwanju Institute for Science and Technology, South Korea). Pax7 was detected using a 1:500 dilution of rabbit anti-Pax7-antibody (AVIVA Systems Biology, San Diego, CA, USA) overnight. Myogenin was visualized using a 1:500 rabbit polyclonal antibody (Santa Cruz Biotechnology, M-225, SC-576) overnight. All primary antibodies were detected using a goat anti-rabbit-IgG secondary antibody (1:5000, Li-Cor Biosciences) for 1 h. Prior to blocking, all immunoblots were treated with Swift Membrane Stain (G. Biosciences, St. Louis, MO, USA) to normalize for sample loading. Band intensities for all antibodies were determined using ImageJ software and normalized to bands visualized using Swift Membrane Stain.
All statistical analysis was performed using GraphPad Prism 5 software. Averaged data are reported as the mean ± the standard error of the mean (SEM). Comparison for two groups was performed using a Students t-test and between multiple groups using Bonferroni post-test with two-way ANOVA on ranks for non-parametric data. For non-paired tests such as fiber size, the Students unpaired t-test with Welsh's correction and between groups Kruskal–Wallis test or Dunn's multi-comparative test. P < 0.05 was considered statistically significant.
This study was supported by grants from Cure CMD, Struggle Against Muscular Dystrophy (SAM), NIH/NIAMS (R01AR053697 to D.J.B. and R43AR057594 to Prothelia, Inc., Milford, MA, USA, to B.L.H.) and NIH/NIGM (8 P20 GM103440: NV INBRE undergraduate student award to P.M.).
The authors thank Drs Paul Martin and Eva Engvall for the dyW−/− mice and Senny Wong, Katie Flynn and Chelsea Lamb for technical assistance.
Conflict of Interest statement: The University of Nevada, Reno, has been issued a patent on the therapeutic use of laminin, laminin derivatives, and their compositions. The patent inventors are Dean J. Burkin and Jachinta E. Rooney. The University of Nevada, Reno has licensed this technology to Prothelia Inc., and has an equity share in this company. B.L.H. is the Chief Scientific Officer of Prothelia and has an equity share in Prothelia.