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Endocrinology, doi:10.1210/en.2004-1594
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Endocrinology Vol. 146, No. 4 1789-1797
Copyright © 2005 by The Endocrine Society

Insulin-Like Growth Factor-I Gene Transfer by Electroporation Prevents Skeletal Muscle Atrophy in Glucocorticoid-Treated Rats

O. Schakman, H. Gilson, V. de Coninck, P. Lause, J. Verniers, X. Havaux, J. M. Ketelslegers and J. P. Thissen

Unité de Diabétologie et Nutrition et Unité de Pathologie Cardio-Vasculaire, Université Catholique de Louvain, B-1200 Brussels, Belgium

Address all correspondence and requests for reprints to: Jean-Paul Thissen, Unité de Diabétologie et Nutrition, Université Catholique de Louvain, 54 avenue Hippocrate, B-1200 Brussels, Belgium. E-mail: thissen{at}diab.ucl.ac.be.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Catabolic states caused by injury are characterized by a loss of skeletal muscle. The anabolic action of IGF-I on muscle and the reduction of its muscle content in response to injury suggest that restoration of muscle IGF-I content might prevent skeletal muscle loss caused by injury. We investigated whether local overexpression of IGF-I protein by gene transfer could prevent skeletal muscle atrophy induced by glucocorticoids, a crucial mediator of muscle atrophy in catabolic states. Localized overexpression of IGF-I in tibialis anterior (TA) muscle was performed by injection of IGF-I cDNA followed by electroporation 3 d before starting dexamethasone injections (0.1 mg/kg·d sc). A control plasmid was electroporated in the contralateral TA muscle. Dexamethasone induced atrophy of the TA muscle as illustrated by reduction in muscle mass (403 ± 11 vs. 461 ± 19 mg, P < 0.05) and fiber cross-sectional area (1759 ± 131 vs. 2517 ± 93 µm2, P < 0.05). This muscle atrophy was paralleled by a decrease in the IGF-I muscle content (7.2 ± 0.9 vs. 15.7 ± 1.4 ng/g of muscle, P < 0.001). As the result of IGF-I gene transfer, the IGF-I muscle content increased 2-fold (15.8 ± 1.2 vs. 7.2 ± 0.9 ng/g of muscle, P < 0.001). In addition, the muscle mass (437 ± 8 vs. 403 ± 11 mg, P < 0.01) and the fiber cross-sectional area (2269 ± 129 vs. 1759 ± 131 µm2, P < 0.05) were increased in the TA muscle electroporated with IGF-I DNA, compared with the contralateral muscle electroporated with a control plasmid. Our results show therefore that IGF-I gene transfer by electroporation prevents muscle atrophy in glucocorticoid-treated rats. Our observation supports the important role of decreased muscle IGF-I in the muscle atrophy caused by glucocorticoids.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
CATABOLIC STATES SUCH as cancer, sepsis, AIDS, or chronic inflammatory diseases are characterized by an accelerated loss of lean body mass, mainly skeletal muscle. This change in body composition is associated with an increase in morbidity and mortality (1). Despite recent better understanding of the metabolic and molecular derangements leading to muscle wasting, therapy of muscle atrophy has still a poor success rate.

Glucocorticoids play a major role in the induction of muscle atrophy in catabolic conditions (2, 3). Indeed, administration of glucocorticoids induces muscle atrophy mainly due to stimulation of muscle proteolysis (4, 5) and inhibition of protein synthesis (4, 6). Furthermore, circulating glucocorticoids levels are frequently increased in catabolic states (2). Finally, the inhibition of glucocorticoids action by RU 486, an antagonist of the glucocorticoid receptor, markedly reduces the loss of muscle mass encountered in such catabolic states (3, 7, 8). Recent evidence indicates that the loss of muscle mass induced by glucocorticoids might be due to decreased IGF-I levels, an important endocrine and autocrine anabolic growth factor (9). Glucocorticoid treatment indeed decreases skeletal muscle IGF-I expression (10), and systemic administration of IGF-I has been reported to prevent corticosteroid-induced muscle atrophy (6, 11).

Several evidences indicate that IGF-I plays a crucial role in the loss of muscle mass observed in catabolic states. First, autocrine IGF-I production plays a critical role in muscle growth (9, 12). IGF-I plays an anabolic and anticatabolic action in skeletal muscle in vivo and in vitro. Anabolic effects of IGF-I result from the stimulation of protein synthesis (13) as well as proliferation and differentiation of muscle satellite cells (14, 15), whereas its anticatabolic effects result from the inhibition of muscle proteolysis (16) and apoptosis (17). Moreover, circulating and muscle IGF-I levels are frequently reduced in catabolic states (18). Finally, IGF-I gene overexpression prevents loss of muscle in some muscle atrophy models such as aging and muscle dystrophy (19, 20). These evidences therefore suggest that IGF-I might prevent loss of muscle mass in hypercatabolic states

However, use of systemic IGF-I administration is limited by its hypoglycemic (21) and cardiac hypertrophic actions (22, 23). Moreover, local production and thus autocrine action of IGF-I seem to play a more important role in the regulation of muscle mass than systemic and endocrine IGF-I (9). Therefore, to limit loss of muscle mass observed in catabolic states, IGF-I administration must mimic as close as possible the autocrine production of IGF-I.

Taking together these observations, we hypothesized that the local decrease in muscle IGF-I content might be responsible for the muscular atrophy induced by glucocorticoids. The aim of this study was therefore to demonstrate that local IGF-I overexpression by electroporation of IGF-I DNA could prevent muscle atrophy caused by glucocorticoid treatment. To demonstrate our hypothesis, we studied the anticatabolic effect of local IGF-I overexpression on skeletal muscle of rats made catabolic by glucocorticoid treatment. The study was performed on the tibialis anterior (TA) muscle, a skeletal muscle quite homogenous (fibers IIb/d 96%) (24) and highly sensitive to the catabolic effects of glucocorticoids (11, 25, 26). To avoid problems encountered with viral vectors (27, 28, 29), in vivo DNA electrotransfer was used to deliver the IGF-I gene into muscle. This method has been already successfully used in several tissues (30, 31, 32) and especially in skeletal muscle (33, 34, 35, 36).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Expression plasmids and DNA preparation
Plasmid pM1-hIGF-I was constructed by inserting a 770-bp human (h) IGF-I cDNA (a generous gift from Dr. Paul Steenbergh, University of Utrecht, The Netherlands) into the SalI-NotI sites, between the cytomegalovirus promoter and a 3' flanking sequence of the bovine GH polyadenylation signal, in the pM1 expression vector (Roche Molecular Biochemicals, Indianapolis, IN). A pM1-ß-galactosidase (gal) expression vector (Roche Molecular Biochemicals) was used to determine the optimal transfection’s conditions. Plasmids were amplified in Escherichia coli top 10 F' (Invitrogen, Carlsbad, CA) and purified with an EndoFree plasmid giga kit (QIAGEN, Hilden, Germany). Plasmids were stocked at –80 C. The day of injection, 100 µg of plasmid were lyophilized and resuspended in 100 µl NaCl 0.9% solution.

Determination of optimal transfection’s conditions by electroporation in muscle
Animals.
Six-week-old male Wistar rats (150–160 g) provided by Janvier Breeding (Le Genest St-Isle, France) were housed under controlled conditions of lighting (12-h light, 12-h dark cycle) and temperature (22 ± 2 C). Access to animal chow and water was unrestricted. Animals were weighed daily.

DNA injection and electroporation.
Rats were anesthetized with a mixture of 75 mg/kg ketamine (Ketalar, Pfizer, New York, NY) and 15 mg/kg xylazine hydrochloride (Rompun, Bayer, Leverkusen, Germany) administered by ip injection. In a first step, we determined the effects of electroporation on muscle transfer of naked DNA. In this experiment, the two tibial anterior (TA) muscles were injected transcutaneously with 100 µl pM1-ß-gal plasmid solution using an insulin syringe with a 30-gauge needle. One TA muscle was electroporated and the contralateral muscle was not. Transcutaneous pulses were applied by two stainless steel plate electrodes with distance between the two plates of 1 cm. Electrical contact with the leg skin was ensured by shaving each leg and applying conductive gel. Electric pulses with a standard square wave were delivered by an electroporator (PA-4000, four-parameter Pulsagile electroporation system, Cytopulse Science, Inc., Columbia, MD). Ten pulses of 200 V/cm were administered to the muscle with a delivery rate of 1 pulse/sec with each pulse of 20 msec in duration. In a second step, we tested the influence of multiple injection sites on the efficiency of transfection by electroporation. In the TA of one side, 100 µl pM1-ß-gal plasmid solution were injected in one single injection and muscle was electroporated as previously described. In the contralateral TA muscle, the 100-µl pM1-ß-gal plasmid solution was divided into 10 injection sites. Electroporation conditions were as previously described, except that the 10 pulses were replaced by two series of five pulses applied on each half of the muscle to cover most of the muscle volume.

Assessment of ß-gal gene expression.
Rats were killed by decapitation 3 d after DNA injection and electroporation. TA muscles were dissected, weighed, snap frozen in liquid nitrogen, and stored at –80 C. According to manufacturer recommendation, 100 mg TA muscle, previously pestled in liquid nitrogen, were homogenized with Ultraturrax (IKA-Labortechnik, Staufen, Germany) in 1 ml reporter lysis buffer (Promega, Madison, WI) and centrifuged 10 min at 10,000 rpm (Sorvall SS-34, GOFFIN-MEYVIS, Overijse, Belgium). Supernatant (150 µl) was then mixed and incubated 30 min with 150 µl of reaction buffer (Promega). The reaction was stopped with 500 µl of carbonate de lithium (1 M), and the absorbance was measured with a spectrophotometer at 420 nm. Muscle protein content was determined by using Bradford’s protein assay (Bio-Rad Laboratories, Munich, Germany). Results were expressed in milliunits of ß-gal per gram of muscle protein. The variation coefficient was calculated as SD/mean of five measurements of ß-gal activity determined in the same electroporation conditions.

Histochemical staining for ß-gal activity.
Rats were killed by decapitation 3 d after DNA injection and electroporation. TA muscles were dissected and 0.5-cm-thick transverse slices were made in the middle belly of the muscle. Five-micrometer-thick sections were obtained with a cryotome (Micron HM500, Micron International, Walldorf, Germany, T = –26 C) and directly applied onto Superfrost Plus slides (Menzel Gläser, Braunschweig, Germany). Sections were dried [15 min at room temperature] and then stored at –80 C until use. To detect ß-gal activity, cryosections were successively air dried (10 min, 37 C), fixed in formaldehyde (10 min at room temperature), washed under tap water, rinsed in PBS (10 min at room temperature), and finally incubated (30 min, 37 C) with a commercial solution of X-gal (ß-gal staining set, ref. 1828673, Roche Diagnostic, Mannheim, Germany).

Evaluation of the anticatabolic effect of IGF-I DNA transfection in glucocorticoid-treated rats
Animals.
Six-week-old male Wistar rats (190–215 g, Janvier Breeding) were used to assess the muscle anticatabolic action of transfected IGF-I in rats treated with dexamethasone. The animals were housed under controlled conditions of lighting (12-h light, 12-h dark cycle) and temperature (22 ± 2 C.).

Experimental design.
Thirty rats were divided into three groups: ad libitum (n = 6), pair fed (n = 12), and dexamethasone (n = 12). The ad libitum and pair-fed groups received daily sc injections of saline solution (vehicle, 0.9% NaCl). The glucocorticoid-treated group received daily sc injections of saline solution of dexamethasone (10 µg per 100 g body weight, Aacidexam, Organon, Oss, The Netherlands). Because dexamethasone treatment is known to decrease food intake, the pair-fed group received the same amount of food as that consumed by the dexamethasone-treated group during the previous day. The ad libitum and dexamethasone-treated animals were allowed free access to chows and water. Animals were weighed and food intake measured every day. Animals were killed after 7 d of dexamethasone treatment. For biochemical analysis, TA muscles were removed, weighed, deep frozen in liquid nitrogen, and stored at –80 C until further analysis. For histological analysis, TA muscles were dissected, and a transversal slice of 0.5 cm thickness was made in the middle belly of the muscle. This slice was embedded in Tissue Tek (Sakura, Zoeterwoude, The Netherlands), frozen in precooled isopentane, and stored at –80 C until further analysis.

DNA injection and electroporation.
Three days before dexamethasone treatment, each rat was anesthetized with a ketamine/xylazine hydrochloride mixture, as previously described. In the ad libitum group, the two TA muscles were injected with 10 µl pM1 plasmid solution (1 µg/µl) in 10 different sites (total volume per muscle = 100 µl). In the two other groups, 10 µl pM1-hIGF-I plasmid solution (1 µg/µl) were injected in 10 different sites in one TA muscle (total volume = 100 µl). The contralateral TA muscle was injected, in the same manner, with 100 µl pM1 plasmid solution. Both muscles were electroporated using the electroporation conditions previously validated.

Muscle protein extraction.
Briefly, 100 mg whole TA muscle, previously pestled in liquid nitrogen, were homogenized with Ultraturrax (IKA-Labortechnik) in 1 ml ice-cold lysis buffer [10 mM Tris/HCl (pH7.5), 5 mM EDTA, 0.5% Nonidet P-40, 1 mM phenylmethylsulfonylfluoride, 5 µg/ml aprotinin, 2 µg/ml leupeptin]. Homogenates were centrifuged 10 min at 10,000 rpm (Sorvall SS-34 rotor) to pellet myofibrillar proteins. Myofibrillar proteins were resuspended in 8 M urea/50 mM Tris/HCl (pH 7.5). The supernatant contained soluble proteins. Myofibrillar and soluble muscle protein contents were determined by using Bradford’s protein assay (Bio-Rad Laboratories).

Muscle and circulating IGF-I extraction.
Five hundred microliters of supernatant, containing soluble proteins, were mixed and incubated 2 h on ice with 500 µl acetic acid (2 M). The resulting homogenate was lyophilized and resuspended in the RIA buffer [0.03 M NaH2PO4, 0.02% protamine, 0.1 M EDTA, 0.05% Tween 20, 0.02% azide (pH 7.5)]. Serum IGF-I was extracted as previously described (37, 38).

IGF-I RIA.
The muscle and circulating IGF-I concentrations were measured by specific RIA (37, 38). Briefly, 200 µl muscle extract or 50 µl serum extract (or standard), IGF-I antiserum, and 125I-IGF-I (10,000 cpm) were pipetted into duplicate RIA tubes. The rabbit anti-IGF-I antiserum (UB-487, a kind gift from Dr. L. E. Underwood, University of North Carolina, Chapel Hill, NC) was used at a dilution of 1:40,000 for muscle IGF-I and at a dilution of 1:15,000 for serum IGF-I. The recovery of recombinant IGF-I added to the crude homogenate was 20.7 ± 0.6% (mean ± SEM, n = 3). The results are expressed as nanograms of IGF-I per gram of muscle and are not corrected for recovery.

RNA extraction and analysis by real-time quantitative (RTQ)-PCR.
Total RNA was isolated from the TA muscle using TRIzol reagent as described by the manufacturer. Recovery was 1 µg/mg TA. Reverse transcription using 1 µg total RNA was done as previously described (39). Specific primers (Table 1Go) were determined using the Primer Express software (Applied Biosystems, Foster City, CA). Accession number for the sequences used were AH002176 (rat IGF-I), S85346 (hIGF-I), and AF106860 [glyceraldehyde-3-phosphate dehydrogenase (GAPDH)] used as reporter gene. Primers were tested to avoid primer dimers, self-priming formation, or unspecific amplification. Human and rat IGF-I primers were tested to demonstrate the absence of cross-reaction with the heterologous IGF-I cDNA sequence. Primers were designed to have standardized optimal PCR conditions.


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  		TABLE 1. Oligonucleotide sequences and variation coefficient of RTQ-PCR

 
RTQ-PCR was carried out using following cycle parameters: 10 min at 95 C, followed by 40 cycles of 1 min at 60 C and 15 sec at 95 C. For each gene, RTQ-PCR was conducted in duplicate with 25 µl reaction volume of 1 ng cDNA. To ensure the quality of the measurements, each plate included for each gene a negative control and a positive control. The threshold cycle (Ct) from a positive sample was used to calculate the intra- and interassay coefficient of variation (CV). For each gene, the intraassay CVs were calculated as SD/mean of five Ct determined for a same sample on the same plate with the same mix. The interassay CVs were calculated as SD/mean of the Ct determined on five different plates and with different mixes. The intra- and interassay CVs obtained for the different genes studied were comprised between 1 and 3% (Table 1Go). Results were expressed using the comparative Ct method as described in the user bulletin 2 (Applied Biosystems). Briefly, the {Delta}Ct values were calculated in every sample for each gene of interest as followed: Ctgene of interest – Ctreporter gene, with GAPDH as the reporter gene. The calculation of the relative changes in the expression level of one specific gene ({Delta}{Delta}Ct) was performed by subtraction of the {Delta}Ct from the control group to the corresponding {Delta}Ct from the treated groups (39, 40) The values and ranges given in different figures were determined as followed: 2–{Delta}{Delta}Ct with {Delta}{Delta}Ct + SEM and {Delta}{Delta}Ct – SEM, where SEM is the SE of mean of the {Delta}{Delta}Ct value (user bulletin 2, Applied Biosystems).

Histological analysis of muscle.
Cryostat serial sections (10 µm thick) were obtained and mounted onto glass slides. The sections were fixed 2 min with acetone for hematoxylin-eosin staining. Muscle and fiber cross-sectional areas (CSAs) were measured with a microscope (Leitz Wetzlar, Brussels, Belgium) coupled to an image analyzer system (Kontron image analysis division, MOP-Videoplan, Eching, Germany). To evaluate muscle fiber CSA, five or six fields were randomly chosen, and 200 fibers per muscle were counted.

Statistical analysis
Results are presented as means ± SEM. Statistical analyses were performed using a one-way ANOVA followed by a Newman-Keuls multiple comparison test to compare muscles from different animals undergoing different experimental conditions (ad libitum-pair fed-dexamethasone) or a paired t test to compare muscles undergoing different experimental conditions within the same animal (pM1-hIGF-I). Fiber CSA distribution statistical analysis was performed using {chi}2 Pearson test. Statistical significance was set at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Optimal transfection conditions
With electroporation (10 pulses of 200 V/cm administered at 1 pulse/sec with each pulse of 20 msec in duration), the ß-gal expression in the ß-gal DNA-injected TA muscle was increased about 30-fold in comparison with nonelectroporated ß-gal reporter plasmid injected contralateral TA muscle (66 ± 17 vs. 2 ± 1 mU/mg protein, P < 0.001, n = 5). When the number of injection sites was increased (10 µg pM1-ßgal plasmid injected in 10 injection sites) and two series of five pulses were applied on each half of the muscle, the ß-gal expression in the TA muscle was increased 6.3-fold, compared with contralateral TA muscle injected with a single injection of ß-gal reporter plasmid and electroporated with one series of 10 pulses (360 ± 46 vs. 58 ± 24 mU/mg protein, P < 0.01, n = 5) (Table 2Go). Under these optimal conditions of DNA injection (10 µg plasmid injected in 10 injection sites) and electroporation (two series of five pulses on each half of the muscle), histological analysis of TA muscles injected with ß-gal reporter plasmid showed an average of 37 ± 8% (mean ± SEM, n = 8) of the fibers expressing ß-gal DNA 3 d after DNA injection and electroporation (Fig. 1Go). These transfection conditions were therefore used to overexpress the IGF-I gene in the muscle of rats.


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  		TABLE 2. Summary of optimal transfection conditions using the pM1-ß-GAL reporter plasmid

 


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  		FIG. 1. Histochemical staining of ß-gal gene electrotransferred TA muscle. TA muscles were analyzed 3 d after DNA injection and electroporation as described in Materials and Methods.

 
Anticatabolic effects of transfected IGF-I in muscle in dexamethasone-treated rats.
Animal weight and food intake.
During the 7-d treatment, the body weight gain was lower in animals treated with dexamethasone (14 ± 2 g) than in the ad libitum rats (64 ± 2 g, P < 0.001) and even in the pair-fed rats (28 ± 2 g, P < 0.001) (Fig. 2AGo). Compared with the ad libitum group, the mean daily food intake was markedly decreased in animals treated with dexamethasone (28.2 ± 0.7 vs. 22.7 ± 0.6 g, P < 0.001). According to our experimental design, the mean daily food intake of the pair-fed group was also reduced at 22.7 g (Fig. 2BGo).



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  		FIG. 2. Dexamethasone treatment reduced body weight gain and food intake. A, Evolution of body weight of the ad libitum (solid line), pair-fed (dotted line), and dexamethasone-treated rats (dashed line). Three days after DNA injection and electroporation (a), rats were treated with dexamethasone (0.1 mg/kg·d) (b) during 7 d and killed (c). B, Daily mean food intake (grams per day) of the three experimental groups. Results are expressed as mean ± SEM. ***, P < 0.001 vs. ad libitum. °°, P < 0.01 vs. pair-fed animals. Dexa, Dexamethasone.

 
IGF-I muscle content.
After 7 d of dexamethasone injections, the IGF-I muscle content was decreased by about 50% in the TA muscle electroporated with the pM1 plasmid, compared with muscle of ad libitum animals (7.2 ± 0.9 vs. 15.7 ± 1.4 ng/g of muscle, P < 0.001, n = 6). Food intake restriction imposed in the pair-fed group was also associated with a significant, albeit smaller (30%), fall in muscle IGF-I content (11.1 ± 0.8 vs. 15.7 ± 1.4 ng/g of muscle, P < 0.05, n = 6). However, in the muscle electroporated with the IGF-I plasmid, the IGF-I muscle content was 2-fold increased by comparison with the contralateral muscle transfected with the pM1 in both the pair-fed (20.7 ± 1.9 vs. 11.1 ± 0.8 ng/g of muscle, P < 0.001, n = 6) and dexamethasone-treated animals (15.8 ± 1.2 vs. 7.2 ± 0.9 ng/g of muscle, P < 0.001, n = 6). Transfection of IGF-I gene was therefore able to completely prevent the decline of the muscle IGF-I content caused by dexamethasone (Fig. 3AGo).



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  		FIG. 3. IGF-I gene electrotransfer restores IGF-I muscle content and increases hIGF-I mRNA in dexamethasone-treated rats. A, TA muscles were injected and electroporated with a total of 100 µg pM1 plasmid at a concentration of 1 µg/µl (white box). In pair-fed and dexamethasone-treated rats, contralateral TA muscles were injected and electroporated with 100 µg pM1-hIGF-I plasmid at a concentration of 1 µg/µl (black box). B, Dexamethasone (Dexa) injection reduced expression of rat IGF-I mRNA (gray box), whereas IGF-I gene electrotransfer increased hIGF-I mRNA (lined box) by comparison with ad libitum control rats. Both muscle IGF-I protein and muscle IGF-I mRNA content were determined 10 d after IGF-I DNA injection and electroporation according to the experimental design described in Materials and Methods. Results are expressed as mean ± SEM. A, ***, P < 0.001 vs. contralateral muscle; °, P < 0.05, °°°, P < 0.001 vs. ad libitum animals. B, ***, P < 0.001 vs. rat IGF-I mRNA in the same muscle; °, P < 0.05 vs. rat IGF-I mRNA in the muscle of ad libitum animals. ND, Not detectable.

 
IGF-I mRNA muscle levels.
Muscle endogenous IGF-I mRNA levels were significantly reduced in the animals treated with dexamethasone (44%, P < 0.05, n = 6) in comparison with ad libitum rats. In contrast, food restriction did not affect rat IGF-I mRNA expression. Using species-specific primers, hIGF-I mRNA was not detected in the TA muscle electroporated with pM1 plasmid. However, in the IGF-I plasmid-electroporated TA muscle, ectopic hIGF-I mRNA was abundantly present. In these muscles, the level of hIGF-I mRNA was even higher than that of endogenous rat IGF-I mRNA (5-fold in the pair-fed and dexamethasone-treated groups, P < 0.001; n = 6) (Fig. 3BGo).

Circulating IGF-I concentrations
Serum IGF-I concentrations did not change significantly as the result of dexamethasone treatment (992 ± 39 ng/ml serum, P > 0.05, n = 11) or food restriction (898 ± 41 ng/ml serum, P > 0.05, n = 12), compared with the levels observed in ad libitum animals (960 ± 53 ng/ml serum, n = 6).

Muscle weight
The muscle weight of the TA electroporated with pM1 plasmid was significantly (13%) lower in dexamethasone-treated rats than in ad libitum rats (dexamethasone: 403 ± 11 mg, n = 5 vs. ad libitum: 461 ± 19 mg, n = 6; P < 0.05). IGF-I transfection increased by about 8% the TA muscle weight in dexamethasone-treated animals (TA electroporated with pM1-hIGF-I: 437 ± 8 vs. TA electroporated with pM1: 403 ± 11 mg, P < 0.05, n = 5). Therefore, by comparison with ad libitum rats, the loss of muscle mass due to the dexamethasone injections was partially prevented in the TA muscle transfected with IGF-I gene (ad libitum with pM1: 461 ± 19 mg vs. dexamethasone with pM1-hIGF-I: 437 ± 8 mg, ns). There was no significant effect of IGF-I overexpression on the muscle weight in the pair-fed animals (Fig. 4AGo).



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  		FIG. 4. IGF-I gene electrotransfer prevents muscle atrophy caused by dexamethasone treatment. TA muscles were injected and electroporated with a total of 100 µg pM1 plasmid at a concentration of 1 µg/µl (white box). In pair-fed and dexamethasone-treated-rats (Dexa), contralateral TA muscles were injected and electroporated with pM1-hIGF-I plasmid at a concentration of 1 µg/µl (black box). Muscle mass (A), total muscle CSA (B) and mean fiber CSA (C) were determined 10 d after DNA injection and electroporation. Results are expressed as mean ± SEM. *, P < 0.05 vs. contralateral muscle; °, P < 0.05, °°, P < 0.01, °°°, P < 0.001 vs. muscle of ad libitum animals.

 
Muscle area and fiber CSA
Dexamethasone decreased by about 11% the TA muscle CSA (ad libitum: 35.8 ± 0.8, n = 6 vs. dexamethasone: 31.7 ± 1.7 mm2, n = 5, ns). This decrease in muscle area was caused by a reduction in the muscle fiber CSA. Indeed, the muscle fiber size was significantly 30% lower in the dexamethasone-treated than the ad libitum animals (dexamethasone: 1759 ± 131 n = 5 vs. ad libitum: 2517 ± 93 µm2, P < 0.001, n = 6). Food restriction also caused a significant reduction by about 19% in muscle fiber size (pair-fed: 2049 ± 111 vs. ad libitum: 2517 ± 93 µm2, P < 0.01, n = 6).

IGF-I gene transfection increased average muscle fiber CSA by about 22% in dexamethasone-injected rats (pM1-hIGF-I: 2269 ± 129 vs. pM1: 1759 ± 131 µm2, P < 0.05, n = 5) and about 9% in pair-fed rats (pM1-hIGF-I: 2250 ± 183 vs. pM1: 2049 ± 111 µm2, P < 0.05, n = 6). In parallel, the average TA muscle CSA tended to increase when IGF-I was overexpressed in the muscle of dexamethasone-injected rats (pM1-hIGF-I: 35.5 ± 1.2 vs. pM1: 31.7 ± 1.7 mm2, n = 5, ns) (Fig. 4Go, B and C). The increase in the average muscle fiber CSA induced by IGF-I transfection was confirmed by the analysis of fiber size distribution showing that the proportion of large fibers was significantly greater in the IGF-I transfected muscle than the pM1 control one in the pair-fed group (P < 0.001) (Fig. 4BGo) and even more markedly in the dexamethasone group (P < 0.001) (Fig. 5CGo). In agreement with the results obtained for the muscle weight, IGF-I overexpression prevented the catabolic effects of the dexamethasone on muscle fiber CSA.



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  		FIG. 5. IGF-I gene electrotransfer prevents decrease of fiber CSA. The proportion of large fibers in the pair-fed (B) and dexamethasone groups (C) was larger in the IGF-I DNA injected TA muscle (black box) than the contralateral pM1 DNA-injected TA muscle (white box). Statistical analysis was performed using {chi}2 Pearson test. ***, P < 0.001 for both B and C.

 
Myofibrillar and soluble muscle protein concentration
The myofibrillar and soluble muscle protein concentrations were not significantly different among all the groups considered in this work. As observed in other muscle atrophy models (41), the muscle atrophy caused by glucocorticoids is characterized by a reduction in muscle mass without any reduction in myofibrillar protein concentration (milligram per gram muscle). Furthermore, the absence of any decrease in muscle protein concentrations in the pM1-hIGF-I groups supports a protein gain and not a water gain (Table 3Go).


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  		TABLE 3. Myofibrillar and soluble muscle protein concentrations in the TA muscle of ad libitum-fed, pair-fed, and dexamethasone-treated rats whose one TA muscle was injected and electroporated with pM1-hIGF-I and the other with pM1

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Our observations show that localized overexpression of IGF-I by gene electrotransfer prevents muscle atrophy in glucocorticoid-treated rats. In this work, high rate of fiber transfection and gene expression was reached by combining multiple injection sites of DNA with electroporation. Human IGF-I gene electrotransfer using this optimized protocol resulted in increased muscle IGF-I mRNA and protein levels associated with the prevention of loss of skeletal muscle mass induced by dexamethasone.

Electrotransfer of naked DNA is a powerful tool for in vivo muscle transfection and a promising avenue for treatment of muscle disorders (42). However, the migration of injected DNA into muscle is very limited (43) and, therefore, distribution over the whole muscle often poor and nonhomogenous. The number of transduced muscle fibers after DNA electroporation has been reported to be around 10% (44). Our observations show that injection of the ß-gal reporter plasmid at multiple sites in the TA muscle followed by electroporation with square wave pulses delivered by plate electrodes increased by several orders of magnitude the reporter gene expression and the number of transfected fibers together with reduction of the interindividual variability (35, 36, 45). In our case, the number of transfected fibers is an important issue, given the fact that IGF-I exerts its anabolic actions toward muscle essentially in an autocrine and paracrine fashion (46). So a marked but restricted expression of IGF-I might not be sufficient to reach a large number of cells and stimulate whole muscle development. With our injection procedure and electric parameters, the average number of ß-gal-positive cells (37%) obtained was higher than with a single injection of naked DNA (10–20%) (34, 36, 47) and equivalent with multiple injection sites (45). Recently Dona et al. (48) obtained by gene electroporation a percentage of transfected muscle cells as high as 80%. However, this high percentage was obtained with an invasive method susceptible to heighten the inflammatory response due to the electroporation. According to our results, the number of transfected cells achieved with our protocol seems largely sufficient to induce autocrine and paracrine effects of IGF-I. The number of transfected fibers is probably less critical in the case of secreted factors with an endocrine action such as erythropoietin (49). Because the TA muscle is almost exclusively composed with type IIb/d fibers (96%) (24), ß-gal-positive cells were for most of them type IIb/d fibers. Whereas electroporation decreased interindividual variability (35), our data indicate that combination of electroporation with multiple injections was able to further reduce this variability.

Glucocorticoid treatment caused a 13% decrease in TA muscle weight. Although this decrease was smaller than that reported by previous studies (15–20%) (10, 11, 50), the difference can be easily explained by the low dose used (0.1 vs. 80, even 5 mg/kg·d) (10, 50) and also by the short period of treatment (7 vs. 28 d) (11) in our study. This decrease in muscle mass resulted from a decrease in mean muscle fiber CSA by 30%, a value comparable with the results of Kanda et al. (5) (–26%) obtained on the extensor digitorum longus and soleus muscle and Fournier et al. (–26%) (11) obtained on diaphragm muscle. We did not investigate the effects of glucocorticoids on muscle fiber phenotype because the TA muscle is almost exclusively composed of type II fibers, a fiber type considered to be more sensitive to the catabolic effects of glucocorticoids than the type I (25, 26). Because the modest food restriction associated with glucocorticoid treatment was sufficient to induce muscle atrophy, the catabolic effects of glucocorticoids in our model partially result from reduced food intake (20%).

Muscle atrophy caused by dexamethasone was associated with a decrease in the IGF-I mRNA and IGF-I peptide muscle content, without any decrease in serum IGF-I concentrations. Nevertheless, some studies reported a decrease in circulating IGF-I concentrations after glucocorticoid administration (10, 51). This decrease was, however, observed only with a large dose of glucocorticoids. Indeed, whereas a high dose of glucocorticoids (80 mg/kg) markedly decreases serum IGF-I (68%) (10), a lower dose (0.3 mg/kg) decreases serum IGF-I only slightly (10%) (51). Compared with these results, the even lower dose of glucocorticoids used in our study (0.1 mg/kg·d) may explain the absence of significant changes in serum IGF-I concentrations. Furthermore, the reduction in serum IGF-I levels observed in the previous studies might be also related to the major reduction in food intake observed in animals treated with high doses of glucocorticoids (52, 53). Indeed, a dose of 80 mg/kg·d of glucocorticoids decreased food intake by about 90% (10), whereas this reduction was only 20% in our study. Finally, it is also possible that high doses of glucocorticoids may inhibit liver IGF-I production independently of the decrease in food intake (51). Altogether these results suggest that the fall in circulating IGF-I levels is not mandatory for glucocorticoids to cause muscle atrophy and therefore indicate that the reduction of the IGF-I muscle content plays a crucial role in the loss of skeletal muscle weight induced by glucocorticoid administration.

The main observation of our study was that overexpression of IGF-I in TA muscle partially prevents muscle atrophy caused by glucocorticoid injections. The increase in muscle mass induced by IGF-I overexpression resulted mainly from increase in the muscle fiber CSA. This increase in TA muscle fiber CSA (+22%) was similar to that found in glucocorticoid-treated animals receiving systemic IGF-I (+15% for the extensor digitorum longus and soleus muscle CSA) (5). In this study, as in our work, IGF-I did not completely prevent muscle atrophy induced by glucocorticoids. Only Fournier et al. (11) totally prevented muscle atrophy caused by glucocorticoids by systemic administration of IGF-I in hamsters.

Interestingly, the muscle anticatabolic effects of electrotransferred IGF-I gene have been recently reported in a hind limb suspension muscle atrophy model (44). In this model, Alzghoul et al. (44) showed that IGF-I overexpression increases gastrocnemius/soleus muscle fiber CSA by about 36% after 7 d of hind limb suspension, compared with the gain of 29% observed after 7 d of glucocorticoid treatment in our work. However, in contrast to our study, which assessed the whole muscle, this study has considered only the transfected fibers and the fibers immediately adjacent to the transfected ones to assess the hypertrophic effect of overexpressed IGF-I (44).

Our results strongly suggest that the anticatabolic action of IGF-I gene transfer results from a complete restoration of IGF-I muscle content in the TA muscle of dexamethasone-treated rats. The doubling of IGF-I muscle content obtained with our electrotransfer protocol is close to the increase in IGF-I observed in muscle IGF-I transgenic mice (20) but lower than the 5-fold increase reached by local infusion of the peptide (54). Only one group has measured the muscle IGF-I content reached after muscle IGF-I gene transfer in a regeneration animal model and showed a modest increase of 30–40% only after 2 wk (55). The results obtained with our local administration of IGF-I contrast with those of Fournier et al. (11), showing that the systemic administration of IGF-I to glucocorticoid-treated animals completely prevents the muscle atrophy without any restoration of the IGF-I levels neither in the muscle nor the circulation.

Changes in muscle IGF-I concentrations were reflected by parallel changes in hIGF-I mRNA levels. Previous observations showed in vitro a down-regulation of IGF-I expression by exogenous IGF-I added to muscle cells in culture (56). However, our observation shows that local increased IGF-I was not associated with a down-regulation of rat IGF-I gene expression. The absence of a negative feedback of IGF-I on its own expression has already been reported in vivo in muscle IGF-I transgenic mice (23).

Some hypotheses can be raised to explain the muscle anticatabolic effect of IGF-I in glucocorticoid-treated rats. First, IGF-I could have inhibited proteolysis induced by glucocorticoids. Indeed, in vivo IGF-I as in vitro IGF-I inhibits total (5, 57, 58, 59) and myofibrillar protein (5, 57) breakdown induced by glucocorticoids. This anticatabolic action of IGF-I could be mediated by the inhibition of the ubiquitin-proteasome-dependent pathway induced by glucocorticoids. Indeed, IGF-I is known to down-regulate glucocorticoid-induced mRNA for several components of the ubiquitin-proteasome-dependent pathway such as ubiquitin, ubiquitin-conjugated enzymes (E214kDa) (50, 60), and especially two ubiquitin ligases, Mafbx/Atrogin-1 and Murf1 (59, 61), crucial in muscle atrophy. IGF-I can also block the lysosomal proteolytic pathway by inhibiting the glucocorticoid-induced cathepsin L (62) and B (58). A second hypothesis is that IGF-I could have increased the protein synthesis inhibited by glucocorticoids. However, the IGF-I anticatabolic action seems unlikely to be due to an increase in protein synthesis because stimulation of protein synthesis by IGF-I seems blunted in presence of glucocorticoids (4).

In conclusion, we have demonstrated that muscle IGF-I gene electrotransfer prevents muscle atrophy caused by glucocorticoids. Our results suggest that the fall in IGF-I muscle content induced by glucocorticoids plays a critical role in the loss of muscle mass observed in this model. Because circulating IGF-I levels are not reduced by glucocorticoids, these observations also suggest that local IGF-I levels play a more critical role in the regulation of muscle mass than endocrine IGF-I. Given that glucocorticoids play a major role in muscle atrophy, our data may be transposable in most catabolic models including sepsis and immobilization in which glucocorticoids are induced. The preservation of muscle mass by IGF-I does not allow to assert that muscle function was improved in parallel. Nevertheless, results obtained in IGF-I transgenic mice indicate that the muscle function was preserved in parallel to the muscle mass (19). Preventing a decrease of IGF-I muscle by local administration of IGF-I genes may provide a clinically relevant avenue for the preservation, attenuation, or reversal of disease-related muscle loss.


   Acknowledgments
 
We gratefully acknowledge Professor Veronique Préat (School of Pharmacy, Université Catholique de Louvain), Professor Jean Lebacq (Departments of Physiology and Pharmacology, Université Catholique de Louvain), and the Pathology Department of Saint-Luc Academic Hospital (Brussels, Belgium), especially Drs. Christine Sempoux and Yves Guiot, for expert technical assistance.


   Footnotes
 
This work was supported by grants from the Fund for Scientific Medical Research (Belgium) and the Fonds Spéciaux de Recherche (Université Catholique de Louvain, Belgium).

First Published Online January 20, 2005

Abbreviations: CSA, Cross-sectional area; Ct, threshold cycle; CV, coefficient of variation; gal, galactosidase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; h, human; RTQ, real-time quantitative; TA, tibialis anterior.

Received December 8, 2004.

Accepted for publication January 7, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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