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,
* Department of Neurosciences, Psychiatry and Anaesthesiology, University of Messina, Messina, Italy;
Department of Experimental Medicine and Pharmacology, University of Messina, Messina, Italy;
Molecular Medicine Laboratory, International Centre for Genetic Engineering and Biotechnology (ICGEB), Trieste, Italy; and
Department of Clinical and Biomedical Science, Faculty of Medicine, University of Trieste, Italy
1Correspondence: Department of Neurosciences, Psychiatry and Anaesthesiology, Unit of Clinical Neurobiology and Neuromuscular Diseases, AOU Policlinico "G. Martino," 98125 Messina, Italy. E-mail: giuseppe.vita{at}unime.it
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: dystrophin gene • muscle necrosis • rAAV • Duchenne muscular dystrophy
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. The disease is due to a mutation in the dystrophin gene and to the consequential lack of this protein, an essential component of the dystrophin-glycoprotein complex that maintains sarcolemmal integrity (2)
. The mechanisms responsible for the morphological hallmarks of the dystrophic process such as necrosis, phagocytosis, infiltration of inflammatory cells, initial efficient regeneration, followed by a decline and secondary fibrosis, have not been definitively identified.
Possible DMD therapeutic approaches are often developed on the genetically homologous murine model, the mdx mouse, despite relevant clinical and morphological differences. The murine model exhibits a milder phenotype, with late muscle weakness, slow disease progression, and similar extensive degeneration and regeneration cycles, but only between 2 and 12 wk of age and not followed by proliferation of connective tissue in limb muscles (3
, 4)
. In DMD, the regeneration process consisting of the activation, migration, and fusion of satellite cells (SC) is not efficient enough, probably due to a limited proliferative capacity of SC to be able to sustain repeated cycles of degeneration and regeneration. Therefore, enhancing the exhaustible regeneration may be a promising therapeutic approach to DMD.
The molecular basis of SC activation has been studied extensively in isolated cultures and to lesser extent in vivo in different experimental models of muscle necrosis (5
, 6)
. Various factors, some of which are secreted by macrophages, can activate SC such as fibroblast growth factor, hepatocyte growth factor, insulin growth factor-I and -II, transforming growth factor-ß, leukemia inhibitory factor, and interleukin-6 (6)
. Recently, several studies highlighted that the vascular endothelial growth factor-A (VEGF-A) also interacts with SC and has a role in promoting regeneration in muscle.
VEGF-A (hereafter, VEGF) is a key regulator of physiological angiogenesis during embryogenesis but has also been implicated in pathological angiogenesis associated with tumors, intraocular neovascular disorders, and other conditions (7)
. Although it was originally described as an endothelial-specific growth factor, recent evidence suggests that the effects of VEGF might extend to a variety of other cell types such as neurons, hepatocytes, osteoblasts, hematopoietic cells, and myoblasts (8)
. The biological effects of the VEGF family members are transduced by three main receptors: VEGFR-1 (Flt-1), VEGFR-2 (KDR, Flk-1), and VEGFR-3 (Flt-4). The VEGF-R2 is the main receptor that mediates the angiogenic and permeabilizing effects of VEGF through the activity on endothelial cell proliferation and migration. Moreover, it was recently demonstrated that VEGFR-2 is also the main mediator of the effect of VEGF on myogenic cells (9)
.
Several studies in vivo and in vitro have highlighted a role of VEGF in promoting muscle regeneration. Studies in vitro showed that, in growing medium, both cultured SC and myoblasts express VEGF, VEGFR-1, and -2 (9
, 10)
. Moreover, VEGF administration in vitro stimulates myoblast migration and survival, protects myogenic cells from apoptosis, and promotes myogenic cell growth (9
, 10)
. In normal muscles, VEGF and its receptors are expressed in vascular structures but not in muscle fibers. After experimental muscle damage, VEGF and its receptors are expressed in SC and in regenerating muscle fibers, suggesting the operation of an autocrine pathway that may promote survival and regeneration of myocytes (9
10
11
12
13)
. In vivo VEGF administration with recombinant adeno-associated viral (rAAV) vectors injected in normal mouse skeletal muscle resulted in the appearance of a notable subset of muscle fibers displaying a central nucleus, a hallmark of muscle regeneration (8)
. Moreover, after experimental muscle damage with ischemia, glycerol, or cardiotoxin, the delivery of AAV-VEGF markedly promoted muscle fiber regeneration with a dose-dependent effect. It is important to note that this proregenerative effect was more evident when VEGF was delivered 5 days after muscle damage, suggesting a direct effect on myogenesis in addition to the well-established proangiogenic activity (8)
.
Furthermore, increased density of SC has been observed adjacent to capillaries (14)
, suggesting a possible role of VEGF in homing circulating progenitor germ cells (PGC) to specific muscle location and/or in regulating the SC pool. Indeed, the PGC can adopt a myogenic fate (15)
, and VEGF is able to recruit PGC (16
, 17)
and might favor muscle regeneration by promoting transdiffentiation or fusion of these cells (18
, 19)
.
Taken together, this evidence suggests that VEGF might increase the efficiency of the regeneration process acting directly on muscle cells and/or through neovascolarization of the necrotic area and the recruitment of PGC from bone marrow. This work hypothesis would also indicate that VEGF administration could have a therapeutic potential in dystrophic muscle. To confirm and clarify this issue, we used a recombinant AAV vector (rAAV) to express VEGF in mdx mice muscles. AAV-VEGF preparations locally injected in muscle allow a long-term expression of VEGF in the tissue and have minimal capacity to spread to other body districts (20)
. The aim of our study was to test the novel hypothesis that VEGF administration may influence the skeletal muscle pathology in mdx mice with respect to the functional, morphological, and biochemical patterns.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 20
, 21)
.
Animals and experimental protocol
Male mdx and wild-type C57BJ/10 (WT) mice were obtained from The Jackson Laboratory (Bar Harbor, ME, USA) and bred in our animal facilities. Mice were housed in plastic cages in a temperature-controlled environment with a 12 h light/dark cycle and free access to food and water. The investigation conformed with the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health (NIH Publication No.85–23, revised 1996). Four-wk-old mdx and WT mice were treated with intramuscular injections of either rAAV-VEGF (n=5, 100 µl, 3x108 vector particles) or rAAV-LacZ (n=5, 3x108 vector particles). Biceps and tibialis anterior (TA) were injected bilaterally with two injections per muscle. After 4 wk, animals were anesthetized with intraperitoneal administration of sodium penthobarbital (80 mg/kg). Blood collected by intracardiac puncture was drawn to analyze creatine kinase (CK) levels; biceps and TA muscles were removed bilaterally, immediately frozen in liquid nitrogen-cooled isopentane, and stored at –80°C for histological and biochemical evaluations.
Animal examinations
Mice were weighed and examined for forelimb strength at baseline and at the end of the experiment. Strength testing consisted of five separate measurements using a grip meter attached to a force transducer that measures peak force generated (Stoelting Co., Wooddale IL, USA). The mouse grabs the trapeze bar as it is pulled backward and the peak pull force in grams is recorded on a display. The three highest measurements for each animal were averaged to give the strength score. We also calculated the degree of fatigue by comparing the first two pulls to the last two pulls. The decrement between pulls 1 + 2 and pulls 4 + 5 gives a measure of fatigue (22)
.
Serum CK evaluation
Blood samples were centrifuged at 6000 g and the serum was stored at –80°C until the day of analysis. Serum CK was evaluated at 37°C using a commercially available kit (Randox Laboratories LTD, Antrim, UK). The results are expressed as U/L.
Western blot analysis
Samples from biceps muscles were homogenized in lysis buffer (1% Triton X-100; 20 mM Tris/HCl, pH 8.0; 137 mM NaCl; 10% glycerol; 5 mM ethylenediamine tetraacetic acid; 1 mM phenylmethylsulfonyl fluoride; 1% aprotinin; 15 µg ml leupeptin). Protein samples (40 µg) were denatured in reducing buffer [62 mM Tris pH 6.8, 10% glycerol, 2% sodium dodecyl sulfate (SDS), 5% ß-mercaptoethanol, 0.003% bromphenol blue] and separated by electrophoresis on SDS (12%) polyacrylamide gel with a kaleidoscope prestained standard proteins (Bio-Rad, Milan, Italy) to achieve a more accurate molecular weight determination. The separated proteins were transferred onto a nitrocellulose membrane using the transfer buffer (39 mM glycine, 48 mm Tris pH 8.3, 20% methanol) at 200 mA for 1 h. The membranes were stained with Ponceau S (0.005% in 1% acetic acid) to confirm equal amounts of protein and were blocked with 5% non-fat dry milk in Tris-buffered saline (TBS) -0.1% Tween for 1 h at room temperature, washed three times for 10 min each in TBS-0.015% Tween, and incubated with rabbit polyclonal antibody against human VEGF-A (code #9543 Abcam, Cambridge, UK) in TBS-0.1% Tween overnight at 4°C. After washing three times for 10 min each in TBS-0.15% Tween, membranes were incubated with peroxidase-conjugated goat anti-rabbit immunoglobulin G (Pierce, Milan, Italy) for 1 h at room temperature. After washing, the membranes were analyzed by the enhanced chemiluminescence system according to the manufacturer’s protocol (Amersham). The protein signals were quantified by scanning densitometry using a bioimage analysis system (Bio-Profil, Celbio). The results from each experimental group were expressed as relative integrated intensity compared with control muscle measured with the same batch. Equal loading of protein was assessed on stripped blots by immunodetection of ß-actin with a rabbit monoclonal antibody (code #4967, Cell Signaling, Celbio, Italy) diluted 1:500 and peroxidase-conjugated goat anti-rabbit immunoglobulin G (Pierce, Milan, Italy) diluted 1:15000. All antibodies are purified by protein A and peptide affinity chromatography.
Histological studies
Transverse cryostat sections 10 µm thick were obtained from the midpoint of the biceps and TA muscle body. The whole muscle cross sections (corresponding to a mean area of 2.10 mm2 in biceps and 2.25 mm2 in TA) stained with hematoxylin-eosin were examined by a blinded observer, using the AxioVision 2.05 image analysis system equipped with an Axiocam camera scanner (Zeiss, Munchen, Germany). The following four areas with patchy distribution were recognized: 1) normal fibers identified by the presence of peripheral nuclei; 2) necrotic fibers identified by pale cytoplasm and phagocytosis; 3) regenerating fibers identified by small size, central nuclei with/without basophilic cytoplasm; 4) centrally nucleated fibers identified by normal size but with central nuclei. The area occupied by normal fibers, necrotic fibers, regenerating fibers, or centrally nucleated fibers was expressed as the ratio of each area divided by the whole muscle cross section as a percentage.
Immunohistochemistry
Immunohistochemical detection of specific antigens was performed on 7 µm-thick transverse cryostat sections from biceps muscles. A rabbit polyclonal antibody against myogenin (1:60; #sc-576, Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) and a mouse monoclonal antibody against developmental myosin heavy chain (dMHC) (1:20; #NCL-MHCd, Novocastra Laboratories Ltd., Newcastle on Tyne, UK) were used to study regeneration (23)
. Myogenin is expressed by differentiating myoblasts (6)
and is considered an "early" marker of regeneration. DMHC was used as a marker of "later" regeneration (24)
. A mouse monoclonal antibody against desmin (1:75; Ylem, Avezzano, Italy) was used to detect activated SC. VEGFR-2 immunolocalization was studied using a rabbit polyclonal antibody anti-Flk-1 (1:100; code #2479, Cell Signaling Technology, Beverly, MA, USA). Capillaries density was investigated with a rat monoclonal antibody against CD31 (PECAM-1) (1:100; #550274, BD Biosciences PharMingen, San Diego, CA, USA). Visualization of antibody staining was achieved using a peroxidase detection system (ABComplex /HRP kit, Dako, Milan, Italy) and the 3,3-diaminobenzidine tetrahydrochloride as chromogen. In the case of dMHC, the AEC substrate chromogen (Zymed Laboratories, San Francisco, CA, USA) was used for color development.
Statistical analysis
Results are expressed as mean ± SD. Statistical comparison between treated and control groups was performed by the 2-tailed Student’s t test on paired samples with the use of InPlotPrism software version 3.0 (GraphPad Software, San Diego, CA, USA). P values <0.05 were considered significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A).
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At baseline, strength and strength normalized to weight were significantly lower in both mdx mice groups (assigned to rAAV-VEGF or to rAAV-LacZ) compared with WT groups (allocated to rAAV-VEGF or to rAAV-LacZ) (P<0.05). However, at the end of the experiment, rAAV-VEGF-treated mdx mice had significantly higher forelimb strength (+19.5%; P<0.05) and strength normalized to weight (+14.9%; P<0.05) compared with control-treated mdx mice (Fig. 1B, C
). In all groups the somatic growth paralleled an increment of strength if compared with baseline values (P<0.05) (Fig. 1B
), but when normalized to weight, only the rAAV-VEGF-treated mdx mice showed significant amelioration in strength (P<0.05) (Fig. 1C
).
At baseline the percentage of fatigue was significantly higher in both mdx mice groups compared with WT groups (P<0.05); there was no significant difference between the two mdx groups. After treatment, we found no difference among rAAV-VEGF and rAAV-LacZ groups. Comparing the data longitudinally, the percentage of fatigue increased in the mdx groups regardless of the treatment (Fig. 1D
).
rAAV-VEGF did not cause any significant change in forelimb strength, strength normalized to body weight, or fatigue of WT animals.
Serum CK evaluation
Low CK levels were observed in WT animals treated with either rAAV-VEGF or rAAV-LacZ (WT+rAAV-VEGF=247±28 U/L, WT+rAAV-LacZ=190±23 U/L). Mdx mice showed increased CK levels (mdx+rAAV-LacZ=2876±79 U/L; P<0.01 vs. WT+rAAV-LacZ). As expected, rAAV-VEGF injections in four muscles did not modify the serum enzyme levels (mdx+rAAV-VEGF=3666±754; mdx+rAAV-LacZ=2876±79 U/L) (Fig. 2
).
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VEGF expression
Western blot analysis performed on whole muscle homogenates revealed that rAAV-VEGF treatment induced a 3-fold increase of overall VEGF expression in WT and mdx animals compared with untreated animals (P<0.05) (Fig. 3
). rAAV-LacZ injections had no effects in both groups.
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Histological studies
Wild-type animals showed a normal architecture of the biceps and TA muscles that was not modified by treatment with rAAV-VEGF and AAV-LacZ (Fig. 4
, Fig. 5
). Biceps and TA muscles from rAAV-LacZ-treated mdx mice both showed necrosis and regeneration.
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Quantitative morphological evaluation of biceps and TA muscles from rAAV-VEGF-treated mdx mice revealed a significant decrease in necrotic area (P<0.05) and an increase in regenerating fibers area (P<0.01) (Fig. 4
, Fig. 5
).
Immunohistochemistry
Immunohistochemical analysis revealed low VEGF-R2 immunoreactivity in WT and rAAV-LacZ mdx muscles (<1 positive fiber/mm2). Increased VEGF-R2 immunoreactivity was observed in rAAV-VEGF-treated mdx muscles (11.2 positive fibers/mm2; P<0.01 vs. mdx+rAAV-LacZ) mainly localized at a sarcolemmal level and only occasionally also present in the cytoplasm (Fig. 6
).
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The number of myogenin-positive SC/ mm2 was higher in rAAV-VEGF-treated than in rAAV-LacZ-treated muscles in WT mice (2.16 vs. 0.1 cells/mm2; P<0.05) as well as in mdx mice (4.16 vs. 1.47 cells/mm2; P<0.05) (Fig. 7
A). Myogenin-positive SC appeared to be activated, as proved by the fact that serial histological sections also stained positive for desmin (Fig. 7B, C
) and were mainly localized close to vessels or in the perifascicular area. The number of myogenin-positive nuclei/mm2 in WT mice was very low in both rAAV-VEGF- and rAAV-LacZ-treated muscles (0.2 nuclei/mm2 in both groups). Notably, rAAV-VEGF-treated muscles in mdx mice showed a significantly higher number of myogenin-positive nuclei than rAAV-LacZ-treated (24.32 vs. 5.58 nuclei/mm2, P<0.001) (Fig. 7C, D).
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The number of dMHC-positive fibers/mm2 in WT mice was very low in both rAAV-VEGF- and rAAV-LacZ-treated muscles (0.6 and 0.5 fibers/mm2, respectively). The rAAV-VEGF-treated muscles of mdx mice showed a remarkable increase in the number of dMHC-positive fibers/mm2 compared with rAAV-LacZ-treated animals (19.8 vs. 5.59 fibers/mm2; P<0.05) (Fig. 7F, G
).
In our experimental conditions, the number of CD31-positive capillaries/fiber was not different between rAAV-VEGF- and rAAV-LacZ-treated WT mice (1.5 and 1 capillaries/fiber, respectively). In contrast, the mdx mice rAAV-VEGF-treated muscles showed a higher number of CD31-positive capillaries/fiber than did those injected with rAAV-LacZ (3.54 vs. 2.36 capillaries/fiber; P<0.05) (Fig. 8
A). When measurements were performed by dividing mdx muscles in normal-sized fibers area and in regenerating fibers area, there was no difference in the former between rAAV-VEGF- and rAAV-LacZ-treated muscles (1.9 and 1.01 capillaries/fiber, respectively), whereas the difference was quite pronounced in the latter (2.41 vs. 1.27 capillaries/fiber; P<0.01) (Fig. 8B-G
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In mdx mice, muscle weakness and necrosis are present at 
4–5 wk of age, then a morphological recovery begins with an apparent stabilization of myopathy (3
, 25
26
27)
. For this reason, we chose to start the study at the beginning of the 5th wk of age and to end it by the end of the 8th wk in order to better verify the effect of VEGF treatment on both functional and morphological patterns.
At the end of the experiment, 4 wk after the injections, we found high expression of VEGF in WT and mdx rAAV-VEGF-treated muscles, confirming the elevated efficiency of the AAV-based transduction system in the absence of a vector-induced inflammatory response (9
, 20
, 28)
; this was particularly evident in a damaged tissue such as the dystrophic muscle.
Previous experimental evidence has demonstrated that in vivo administration of AAV-VEGF exerts a powerful effect on muscle survival and regeneration in different models of muscle necrosis (9)
. With the present work, we substantiate this specific action of VEGF in mdx mice. In our study, the histological findings consisted of a significant decrease in the area occupied by necrosis and an increase in the area occupied by regenerating fibers in rAAV-VEGF-treated vs. AAV-LacZ-treated mdx mice. Moreover, the proregenerative effect was confirmed by immunohistochemical results. rAAV-VEGF treatment increased the number of myogenin-positive activated SC not only in mdx, but also in WT mice, in keeping with the reported proregenerative effect in normal muscle (9)
. rAAV-VEGF treatment also augmented the number of myogenin-positive nuclei of differentiating myoblasts and of dMHC-positive cells compared with rAAV-LacZ muscles.
These morphological findings paralleled a positive effect on functional parameters. At the end of experiment, rAAV-VEGF-treated mdx mice showed a higher level of strength and of strength normalized to weight compared with baseline and AAV-LacZ-treated mdx mice. Although there was no difference between the two groups in terms of fatigue, this is probably due to the different response of this parameter to localized administration.
Recently, Wagatsuma et al. further supported the role of VEGF in muscle regeneration, showing in a model of muscle freeze injury that expression of VEGF and its receptors is coordinated with the process of muscle regeneration in vivo and that the muscle fibers express these molecules at most 5 days after damage (13)
. In that study, VEGF and VEGFR-2 were detected in the cytoplasm and on the cell membranes of some regenerating fibers. Also in agreement with other previous observations in experimental models of muscle necrosis (9
10
11)
, we confirm that, in rAAV-VEGF-treated mdx mice, the level of VEGFR-2 is augmented. This receptor is mainly detectable at the sarcolemmal/subsarcolemmal level and its increased expression parallels improves regeneration.
The biochemical pathway that is triggered after VEGFR-2 activation in muscle cells and is responsible for the direct effect of VEGF on myogenesis is still unclear. However, in endothelial cells both PI3K/Akt and MAP kinase signaling pathways, known to be important for muscle survival and regeneration, are set in motion by the activation of VEGFR-2 (29
, 30)
. In muscle cells, the activation of Akt signaling inhibits apoptosis during differentiation (31
, 32)
and regulates myofiber size (33
, 34)
. Muscle fibers transduced by a constitutively active Akt formed in vivo also produce increased levels of VEGF and show signs of muscle hypertrophy (29)
. Furthermore, MAP kinase signaling leads to the increased expression and activity of the MyoD protein involved in cell differentiation (35)
. It is intriguing that insulin growth factor-1, a powerful promoter of muscle regeneration with well-known beneficial effects in mdx mice (36
, 37)
, also activates PI3K/Akt and the MAP kinase pathways and increases VEGF synthesis in C2C12 cells (29)
. These data further support an important role for VEGF in the regeneration process that needs to be better elucidated, with particular attention to rAAV-VEGF-treated mdx mice.
Another factor that likely helps to improve recovery after muscle damage or cooperates in the dystrophic process is the well-known proangiogenic activity of VEGF. In the present study we injected a relatively low dosage of rAAV-VEGF, since high local concentrations of VEGF might be detrimental for the regeneration process by generating excessively disorganized vascular growth. High local concentrations of VEGF indeed determine the formation of a highly permeable vascular structure and the generation of arterovenous shunts that might be detrimental for functional muscle perfusion (20
, 38)
. At lower VEGF dosages, this negative vascular effect is not observable and muscle regeneration appears to predominate (M. Giacca, unpublished observation). Longer term studies are required to determine whether the continuous expression of VEGF will eventually lead to further progression of the angiogenic process or to maturation of the neoformed vessels.
In our study, rAAV-VEGF treatment induced a significant angiogenic effect only in mdx mice, particularly in the area of regenerating fibers, with an increase of > 100% of the number of capillaries per fiber. This finding confirms previous observations on VEGF-mediated neoangiogenesis in damaged muscle tissues already detectable 1 (9)
and 3 months after AAV injection (20)
.
VEGF-induced muscle neovascolarization can have different beneficial effects on dystrophic tissue by 1) promoting macrophage recruitment and removal of cellular debris; 2) increasing release and circulation of factors secreted by mononuclear cells and activating myogenic cells (39
, 40)
; and 3) increasing the recruitment of bone marrow-derived mononuclear cells, which in turn release factors that activate the myogenic process. Recent evidence indicates that VEGF acts a powerful recruiter of bone marrow-derived mononuclear cells (41
, 42)
. These cells are not directly incorporated into the newly formed vasculature and most likely act in a paracrine manner on the angiogenetic process. It needs to be established whether the same cells might directly adopt a myogenic fate in the muscle or might contribute to the myogenic process in a paracrine fashion (43)
.
The present study shows that treatment with AAV-VEGF ameliorates the dystropathology and functional parameters in mdx mice. The main effects seem to be proregenerative and proangiogenic. Further studies are required to evaluate molecular pathways as well as long-term benefits of rAAV-VEGF treatment, as it opens new therapeutic prospectives in DMD and other types of muscular dystrophies. With reference to the therapeutic potential of these observations, AAV vectors have already entered human clinical experimentation for treatment of a number of genetic diseases due to their excellent profile of safety and efficiency. The possibility of inserting tissue-specific or tissue-inducible promoters will offer additional advantages for the therapeutic use of these vectors. Moreover, the recent availability of new AAV vector serotypes (such as AAV9) that determine a widespread muscle transduction after a single systemic injection in mice (44)
will likely allow us to overcome the major hurdle of broader gene transfer in muscle.
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ACKNOWLEDGMENTS |
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Received for publication March 7, 2007. Accepted for publication May 10, 2007.
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