J. Anim. Sci. 2005. 83:2330-2338
© 2005 American Society of Animal Science
ANIMAL GROWTH, PHYSIOLOGY, AND REPRODUCTION |
Ontogeny of factors associated with proliferation and differentiation of muscle in the ovine fetus1,2
A. J. Fahey,
J. M. Brameld,
T. Parr and
P. J. Buttery3
Division of Nutritional Sciences, School of Biosciences, Sutton Bonington Campus, The University of Nottingham, Leicestershire LE12 5RD, U.K.
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Abstract
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The number of muscle fibers within a muscle has been found to be of high importance for the growth potential of an animal, and this number is set during fetal development. The objective of this study was to identify the ontogeny of muscle cell differentiation and fiber formation by observing the changes in expression of factors known to influence myoblast proliferation and differentiation. Twenty-one Swaledale x Leicester Blue Face ewes carrying twins were allotted to this trial. From d 40 of gestation, three ewes were killed every 15 d until term. At each time point, the fetuses were located, removed, and total muscle from both hind limbs was dissected from each fetus and snap frozen in liquid N2. Ribonuclease protection assays were used to quantify transcripts for IGF-I, IGF-II, GH receptor (GHR), and myostatin genes in the muscle samples, whereas quantitative real-time PCR was used to quantify myogenin transcripts. Histological sections also were taken from the fetal muscle samples and observed for evidence of muscle differentiation resulting in fiber formation. The abundance of mRNA for ovine IGF-II and ovine myogenin peaked at d 85 of gestation (P < 0.001). The abundance of ovine IGF-I transcripts peaked at d 100 of gestation, whereas the abundance of ovine GHR mRNA increased throughout gestation (P < 0.05). No change (P = 0.87) in the abundance of myostatin mRNA was observed. The histological sections from the muscle samples demonstrated a clear change in the appearance of the muscle tissue at each time period. Major fiber formation was observed around d 85. The results obtained from the analysis of gene expression and the histological sections suggest that the majority of muscle differentiation and fiber formation takes place around d 85, with myoblast proliferation mainly occurring before this time. It may be possible to manipulate the number of muscle fibers formed by targeting treatments during this proliferation stage immediately before the period of major fiber formation.
Key Words: Muscle Differentiation • Myogenin • Sheep
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Introduction
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In farm animals, muscle fiber number is important for meat content and quality (Rehfeldt et al., 1999
). A change in the number of fibers that form during myogenesis could have a profound affect on the total muscle mass of the mature animal (Bass et al., 2000) and long-term growth potential of the animal. The ability to manipulate fetal muscle fiber number could have important consequences on the postnatal growth of the animal. Once muscle fibers have formed, manipulation of their numbers is thought to be no longer possible. The onset of proliferation and/or differentiation in my-oblasts is controlled by cellular signals. Disruption of IGF-I or -II gene expression has a significant detrimental effect on overall growth, including muscle growth (De Chiana et al., 1990; Liu et al., 1993). The introduction of IGF-II antisense oligonucleotides partially blocked differentiation of muscle cell lines in culture, suggesting a critical role for IGF in myogenic differentiation (Florini et al., 1991b). The positive effects of the IGF on muscle differentiation seem to be via their effects on myogenin gene expression (Florini et al., 1991a). The appearance of myogenin coincides with the onset of differentiation, the lack of which leads to perinatal death in mice due to deficiency of differentiated muscle fibers (Hasty et al., 1993). Myostatin is an inhibitor of myoblast proliferation and differentiation. Joulia et al. (2003) suggested that myogenin is a target of endogenous myostatin, and that the inhibitory influence of myostatin on myoblast differentiation is mediated by its negative control on myogenin expression. This study determined the ontogeny of muscle cell differentiation and major fiber formation by quantifying the mRNA abundance of the above factors that are known to influence myoblast proliferation and/or differentiation. Sections of ovine fetal muscle were taken at time points throughout gestation and examined for evidence of muscle fiber formation.
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Materials and Methods
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Animals
All procedures were performed under the 1986 U.K. Animals (Scientific Procedures) Act and were carried out within the normal seasonal breeding cycle of the ewe. Twenty-one pregnant Swaledale x Leicester Blue Face ewes that were carrying twins were used. The ewes were out to grass until late December, at which point they were group housed and fed a normal commercial diet of grass nuts (cut, dried, and pelleted rye grass) and concentrates, with the amount of concentrates being increased from 0.1 to 0.5 kg (as-fed basis) from mid to late gestation. The ewes were mated naturally with a Charolais ram. Mating was monitored using a raddle placed on the chest of the ram, to which marker paint was applied daily before running with the ewes. The tail area of the ewes was checked every 2 d for fresh paint. Day 0 of gestation was taken as the first day at which the ewes were observed to have an obvious raddle mark. On d 40 of gestation, the ewes were scanned using an ultrasound scanner so that those carrying twins could be identified.
Sample Collection
Three ewes were sacrificed at each time point (d 40, 55, 70, 85, 100, 115, and 130 of gestation). The ewes were weighed and then injected with an overdose of sodium pentobarbitone (130 ng/kg of BW; Vetoquinol UK Ltd., Bicester, U.K.). The abdomens of the ewes were opened, and the uterus was removed intact. The fetuses were located and removed and weighed. Total muscle from both hind limbs was dissected from each fetus (n = 6 at each time point of gestation) and snap frozen in liquid N2. This same procedure was carried out at each time point throughout gestation. The older fetuses (
d 70) were also given an overdose of sodium pentobarbitone via cardiac puncture. In very early gestation, it was impossible to dissect individual hind limb muscles; thus, at each time point, the total muscle available from both hind limbs was dissected and therefore all of the muscle samples (taken at each time point) were of mixed muscle types.
Histology Samples
sThe samples were stored at –80°C before sectioning. The muscle sample was frozen onto a chuck of a cryostat microtome using optimal cutting temperature compound (IMEB, San Marcos, CA). The sample was then allowed to warm to –20°C, which was the temperature of the cryostat chamber. The muscle sample was trimmed with a few rough cuts until an even surface of the muscle was produced. Sections of 20-µm thickness were cut from the smoothed surface and placed onto a microscope slide. The orientation of the muscle was determined by staining with hematoxylin and eosin. The microscope slide holding the muscle section was dipped in hematoxylin (Sigma-Aldrich, Dorset, U.K.) for 30 s, and then the section was gently washed with water for 1 min before counterstaining with 1% eosin (Sigma-Aldrich) for 30 s. The section was then dehydrated using increasing amounts of alcohol. First, the muscle section was dipped in 70% ethanol for 10 s, followed by 95% ethanol for 10 s, and then absolute ethanol for 10 s. The section was then placed in histoclear (II), a histological clearing agent (National Diagnostics, Atlanta, GA), for 30 s before viewing the muscle section under the microscope. From each fetus at each time point throughout gestation, eight sections were taken from different regions within each muscle sample. The stained sections were examined for evidence of muscle fiber formation. Each of the sections was observed so that a representative visual account of the changes in muscle fiber formation during gestation could be obtained. To complement this evaluation of muscle fiber formation, gene transcription analysis relating to muscle differentiation also was carried out on the muscle samples.
Ribonuclease Protection Assay
Ribonuclease protection assays were used to analyze the gene transcription in ovine muscle samples. The abundance of mRNA for IGF-I, IGF-II, GH receptor (GHR), and myostatin was expressed as units per total RNA extracted. Total RNA was extracted from muscle samples that had been stored at –80°C, using an acidified phenol guanidine thiocynate method adapted from Chomcynski and Sacchi (1987). The RNA concentration was quantified by measuring the absorbance at 260 nm (Ultrospec III, Amersham Pharmacia Biotec, Buckinghamshire, U.K.). The ovine-specific riboprobes for IGF-I, IGF-II, and GHR were as previously described (Brameld et al., 2000). For myostatin, the published ovine DNA sequence (Accession No. AF019622; McPherron and Lee, 1997) was used to design specific oligonucleotide primers to generate DNA by PCR, which was then cloned into the pGEM expression vector (Promega, Southampton, U.K.). The myostatin primers used were as follows: forward 5'TCCTTGGAAGACGATGACTAC-CAC 3'; reverse 5' AAGACTCCTAC AACAGTGTTT GTG 3'. Each 32P-labeled riboprobe was generated by in vitro transcription using a DNA template as previously described (Brameld et al., 2000). Hybridization of 40 µg of total RNA with the riboprobes was carried out with the Ambion RPA II kit as specified by the manufacturer (Ambion, Austin, TX). The sample was then denatured as specified by the manufacturer (Ambion), separated on a denaturing urea-polyacrylamide gel, dried, and visualized with phosphoimaging screens (Kodak, Rochester, NY). The amount of target RNA was calculated by measuring the density (intensity x area of band) of the bands produced (using image analyst software; Aida 3.2; Raytest GmbH, Straubenhardt, Germany). The continual changes that occurred due to the status of the animal (i.e., fetal development) presented a problem in finding a factor within the muscle that was constitutively expressed; this is a problem when quantifying the protection assays. A control RNA sample, extracted at the same time as all of the other samples, was routinely used to generate a dilution curve by using increasing quantities and checking that the density increased linearly. If the expected increase in density was not observed, the data from that gel were not used. The same sample also was loaded onto each gel to act as a gel-to-gel standard.
Quantitative Real-Time PCR
Quantitative reverse transcriptase real-time PCR was used to assess myogenin mRNA abundance in ovine muscle samples. A pool of first-strand cDNA was made from 5 µg of skeletal muscle total RNA using random hexamers (Promega) and avian myeloblastosis virus reverse transcriptase (Promega) using conditions as described by the manufacturer in a final volume of 25 µL. Primer Express v. 1.5 software (Applied Biosystems, Foster City, CA) was used to design the primers. The published sequences for bovine (Accession No. AF091714; Oldham et al., 2001), human (AF050501; Tseng et al., 1999) and porcine (SSDNAMYO; Soumillion et al., 1997) myogenin were aligned using the Clus-talW program (www.dur.ac.uk/biological.sciences/index2.htm). A region was chosen where the sequence was highly conserved in these three species, and specific oligonucleotide primers were made. The myogenin primers used were as follows: forward 5' CCTGCCGT G GGCGTGTAAGG 3'; reverse 5' AGATCCTGCGCA GCGCCATC 3'. All real-time PCR was carried out using the SYBR green fluorescence method with universal SYBR green PCR master mix (Applied Biosystems) as specified by the manufacturer. The real-time PCR reactions were carried out in triplicate on an ABI Prism 7700 sequence detection system (Applied Biosystems) using standard default thermal cycling conditions. A pool of ovine muscle cDNA was used to create a standard curve for quantification of the transcripts using a relative standard curve method as described by Applied Biosystems (1997), from which the Ct value (cycle number at which the reporter dye emission intensity rises above background noise) of a particular variant could be converted to nanograms of total RNA equivalent used for first strand synthesis.
Western Blotting
The relative abundance of the desmin protein was measured in whole fetal muscle homogenates using immunoprobing of Western blots. Muscle samples that had been snap-frozen in liquid N2 were finely ground using a pestle and mortar that had been cooled using liquid N2. The fine muscle powder was then homogenized (Polytron PT 3000, Kinematica, Cincinnati, OH) in 10 volumes of buffer (20 mM Tris, 5 mM EDTA, adjusted to pH 7.5 with 1 M HCl), containing 200 µg/mL of 2-(4-aminoethyl)-benzenesulphonyl fluoride, 1 µg/mL of leupeptin, and 1 µg/mL of pepstatin, to protect against proteolytic digestion of the protein samples. The samples were separated on 8% SDS-PAGE (Laemmli, 1970), and then electroblotted onto a nitrocellulose membrane and subsequently probed with the primary mouse desmin monoclonal antibody D1033 (1:300 dilution; Sigma-Aldrich) at room temperature for 45 min. The membranes were then incubated with anti-mouse IgG conjugated with alkaline phosphatase (1:5000; Amersham, Pharmacia Biotech) at room temperature for 30 min and visualized using an enhanced chemiluminescence detection kit (Amersham). The expression of the desmin protein was measured per unit protein of the muscle homogenate. The concentration of total protein in each muscle homogenate was determined using the Amersham PlusOne 2-D Quant kit (Amersham). The density of the desmin band obtained from western blotting was quantified with Multi Analyst image software (Quantity One, BioRad, Hertfordshire, U.K.).
Data Analysis
The levels of IGF-I, IGF-II, GHR, myostatin, myogenin, and desmin at each time point were analyzed by a general linear regression and one-way ANOVA, using the Tukey test as a post-hoc analysis. A general linear model was carried out on the data, which showed that any variation in the results due to the fetuses coming from different ewes was not significant (P = 0.67). Therefore, in this study, n was defined as the number of fetuses not the number of ewes, as each fetus was taken to be independent from other fetuses. The statistical software used was Genstat version 6 for windows (Lawes Agricultural Trust, Hertfordshire, U.K.). Data were considered significant when P < 0.05.
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Results and Discussion
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The main focus of this study was to determine the time during gestation at which the majority of muscle cell differentiation occurred that gave rise to muscle fiber formation. From the histological examination of the fetal muscle samples a clear change in the appearance of the muscle tissue was observed at each time period (Figure 1). The sections in Figure 1 are representative of all the sections taken at each time point. The sections taken on d 40 and 55 showed only fibroblastic cells still undergoing proliferation. On d 70, the muscle tissue was composed of some myotube-like cells; however, it was not until d 85 that major fiber formation and clear organization of the muscle tissue was observed. Although not readily apparent in Figure 1, at d 85 the myotube-like cells were less tubular in shape, which suggested that the myotubes had differentiated into myofibers. From d 100 to 130, there was mainly hypertrophy of the fibers. It was unclear from these sections which were primary and which were secondary fibers. Although areas were selected randomly from serial sections of each sample, it is important to mention that although the results shown are representative for a particular sample of each muscle, it would be misleading to definitely assume that the result is completely representative of the whole muscle. The histological analysis is purely descriptive and provides a visual account of the changes in muscle fiber formation during gestation. This histological examination supplemented the more extensive gene transcription and protein analysis performed on the muscle samples.
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Figure 1. Representative sections of ovine muscle samples taken during gestation at d 40 (A), 55 (B), 70 (C), 85 (D), 100 (E), and 130 (F). Multiple cross sections (20 µm thick) of muscle samples (n = 6) of mixed hind limb muscles were stained using hematoxylin and eosin. Magnification = 200x.
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The majority of peptide growth factors associated with muscle stimulate myoblast proliferation and prevent differentiation (Coolican et al., 1997). In contrast, the various IGF have been observed to stimulate both proliferation and differentiation of myoblasts in vitro. Cell culture experiments have shown that IGF-I and IGF-II are autocrine signals acting during myogenesis that regulate myoblast proliferation and differentiation (Florini et al., 1986; Magri et al., 1994). Work by Musaro and Rosenthal (1999) to determine the role of IGF-I on muscle cell differentiation established IGF-I as an important regulator of muscle cell differentiation through its ability to induce myotube hypertrophy and increase expression of muscle specific genes. In the current study, the abundance of IGF-I mRNA in fetal muscle samples peaked at d 100, with higher levels at d 70, 85, and 100 compared with d 115 and 130 (P < 0.001; Figure 2). This result is consistent with previous work in this area. Our previous cell culture studies (Brameld et al., 1999) showed an increase in IGF-I mRNA with differentiation in fetal sheep myoblasts. Previous in vivo studies have described higher levels of total IGF-I mRNA in fetal sheep muscle at d 84 compared with d 134 (Dickson et al., 1991). Gerrard et al. (1998) investigated the developmental expression and location of IGF-I and IGF-II mRNA and protein in skeletal muscle. The authors stated that skeletal muscle IGF production is regulated in a time-dependent manner in fetal development and arises specifically from developing muscle fibers, and that the change in the muscle IGF-I and II expression coincided with morphological changes in the muscle fiber populations. They concluded that increased expression of IGF-I during the latter stages of gestation was probably related to muscle fiber maturation rather than proliferation (Gerrard et al., 1998). The decreased abundance of IGF-I mRNA after d 100 in the present study may reflect a decrease in IGF-I mRNA once the muscle cells are fully differentiated and innervated.
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Figure 2. Relative abundance of ovine mRNA for IGF-I (A), IGF-II (B), GH receptor (GHR; C), and myostatin (D) in fetal muscle samples on d 40, 55, 70, 85, 100, 115, and 130 of gestation. After performing a ribonuclease protection assay on the total RNA extracted from fetal muscle samples (total hind-limb muscle) at each time point (n = 6) during gestation, samples were separated by polyacrylamide gel electrophoresis and visualized by autoradiography. The amount of target RNA was quantified by measuring the intensity of the bands produced. The abundance of IGF-I mRNA was greater (P < 0.001) on d 70, 85, and 100 than on d 115 and 130. The abundance of IGF-II mRNA was greater (P < 0.001) on d 85 of gestation than on d 40 to 70 and d 100 to 130. Abundance of GHR mRNA increased (P < 0.05) throughout gestation. Abundance of myostatin mRNA did not differ (P = 0.87) among days of gestation. Bars denote mean value (±SEM).
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In porcine fetal myoblasts, IGF-II expression peaks at d 59 after conception and decreases thereafter (Gerrard et al., 1998). This finding is generally consistent with the present study, where the abundance of ovine IGF-II mRNA in the fetal muscle samples peaked at d 85 of gestation (Figure 2
) compared with d 40 to 70 and d 100 to d 130 (P < 0.001). This result agrees with our previous in vitro studies (Brameld et al., 1999), in which IGF II mRNA increased with differentiation in fetal sheep myoblasts. Interestingly, the increase in IGF-II mRNA was observed coincident with the increase in creatine kinase activity, whereas there was a slight delay in the increase in IGF-I mRNA (Brameld et al., 1999). This finding suggests that local IGF-II expression may be more important in regulating differentiation, whereas local IGF-I expression may be a consequence of the differentiation. Indeed, the inhibition of myoblast differentiation by fibroblast growth factor is associated with decreased expression of IGF-II mRNA (Rosenthal et al., 1991). In a review of regulatory factors in the control of muscle development, Dauncey and Gilmour (1996) briefly commented on the levels of IGF-I and II in fetal tissues. They stated that the concentration of IGF-I and II are uniformly low in fetal tissues throughout gestation, except in muscle at the time when secondary fiber formation is taking place.
In the present study, the abundance of mRNA for ovine GHR in the fetal muscle samples increased throughout gestation (P < 0.05; Figure 2). This finding is similar to our previous in vitro studies in differentiating fetal sheep myoblasts, where GHR expression increased with differentiation (Brameld et al., 1999). The expression of GHR has been observed to increase during gestation in other studies. Klempt et al. (1993) found the expression of GHR mRNA in ovine fetal muscle samples to be very low on d 51 of gestation; however, by d 95, the levels of GHR mRNA in the muscle had significantly increased and levels had increased again by d 120. The biological actions of GH on growth and metabolism are initiated by binding of the hormone to its receptor (Isaksson et al., 1985). This interaction with the GH receptor in the liver and other peripheral tissues then initiates expression and secretion of IGF-I (Hynes et al., 1987), giving rise to both direct and indirect effects of GH on growth and metabolism. However, the effect of GH on skeletal muscle is still unclear because in vitro studies have suggested that GH has no direct effects on fetal sheep myoblasts in culture (Harper et al., 1987). Hence, the significance of this increase in GHR mRNA is unknown.
In the present study, no significant change (P = 0.87) in the abundance of myostatin mRNA was observed at the different time points in gestation (Figure 2). Since myostatin was first described by McPherron et al. (1997), much interest has surrounded its expression, as it has been shown to play an important role in muscle development. Natural mutations in the myostatin gene have been shown to be associated with double muscling in cattle (Kambadur et al., 1997), which have a greater number of muscle fibers than normal cattle. Myostatin-null mice show a dramatic and widespread increase in skeletal muscle mass due to an increase in number of muscle fibers (hyperplasia) and thickness of fibers (hypertrophy; Lee and McPherron, 1999). It also plays an important role in fully developed skeletal muscle. Muscle that is undergoing hypertrophy following mechanical stimulation shows an increase in myostatin (Sakuma et al., 2000). Conversely, in regenerating muscle, myostatin levels are decreased (Yamanouchi et al., 2000).
Myostatin, unlike myogenin and the IGF axis, does not have a stimulatory effect on muscle development but is an inhibitor of myoblast proliferation and differentiation. If myostatin is a clear determinant of prenatal muscle growth, then it is perhaps surprising that in the present study, no significant change (P = 0.87) in the abundance of mRNA for myostatin was observed at the different time points in gestation. Other studies have shown myostatin expression very early in gestation (Ji et al., 1998); hence, it is possible that any significant changes observed in myostatin mRNA may have been missed. The first time period in which muscle samples were collected in this trial was d 40, by which time the ewe was already into its second month of gestation. A number of myostatin gene mutations also have been described in cattle that result in varying effects on muscling. The South Devon carries the same myostatin deletion as that observed in the Belgian Blue, but does not have a double-muscling phenotype (Smith et al., 2000). This finding suggests that the lack of myostatin is not the only factor responsible for double muscling. The lack of change in abundance of myostatin transcripts in this study may reflect that with regard to muscle differentiation and fiber formation, myostatin may not be as important as some of the other factors observed.
The results from this study showed a peak in the abundance of ovine myogenin mRNA in the fetal muscle samples at d 85 of gestation (Figure 3) compared with d 40 to 70, (P < 0.001), d 115 to 130 (P < 0.001), and d 100 (P < 0.05). Factors that are expressed around the beginning of differentiation would obviously give an early indication of when differentiation of the muscle occurred. The appearance of myogenin coincides with the onset of myoblast differentiation and fiber formation. Under conditions of growth, myoblasts proliferate and do not express markers of differentiation such as myogenin (Maltin et al., 2001). Myoblast differentiation has been shown to be inhibited in vitro by antisense oligomers that prevent myogenin expression (Florini and Ewton, 1990). Myogenin is a muscle specific transcription factor and has been observed in all myogenic cell lines studied to date. Targeted gene disruption in mice, using homologous recombination to inactivate the myogenin locus, resulted in extreme muscle deficiency due to the inability of individual myoblasts to fuse into muscle fibers (Hasty et al., 1993). Because myogenin expression has been shown to be crucial for myoblast differentiation, it is likely that this peak in myogenin occurs when the major population of the myoblasts is differentiating to form fibers.
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Figure 3. The relative abundance of ovine mRNA for myogenin in fetal muscle samples (total hind-limb muscle) at each time point (n = 6) during gestation, as determined by quantitative reverse transcriptase real-time PCR. A pool of ovine muscle cDNA was used to create a standard curve for quantification of the transcripts using a relative standard curve method as described by Applied Biosystems (1997). Bars denote mean values (±SEM), d 85 compared with d 40 to 70 and d 115 to 130, P < 0.001; d 85 compared with d 100, P < 0.05.
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In the present study, the expression of desmin increased throughout gestation, with an increase (P < 0.001) observed around d 85 (Figure 4). Although the samples continued to show increased expression of des-min, this increase was not significant. Desmin is a major protein of the skeletal muscle intermediate protein system, and plays a critical role in maintaining the registry between adjacent Z-discs. Its importance in myogenesis has been shown in studies using desmin null (knockout) mice, which indicate that normal muscle development depends on the presence of desmin (Smythe et al., 2001). The results indicated that the absence of desmin slightly prolongs myoblast proliferation, resulting in delayed fusion of the myoblasts into muscle fibers. As muscle cells differentiate, an increase in desmin would be expected due to the role of the protein (i.e., maintaining structure). The significant increase of desmin around d 85 probably reflects an increase in the number of fibers formed and hypertrophy of those fibers afterward.
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Figure 4. Relative abundance of ovine desmin protein in fetal muscle samples (total hind-limb muscle). Approximately 40 µg of total protein extracted from fetal muscle samples at each time point (n = 6) during gestation were run on 8% SDS PAGE. After Western blotting, the blots were probed with antidesmin and then visualized with an enhanced chemiluminescence kit. Band intensity was measured using multianalyst software. Bars denote mean value (±SEM), P < 0.001.
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Variation in muscle fiber number is common within different breeds of the same species. In the pig, two thirds of the phenotypic variation in muscle fiber number is due to genetic origin (Rehfeldt et al., 1999). Coefficients of heritability estimated for muscle fiber number range from 0.12 to 0.88 (Rehfeldt et al., 1999), demonstrating that muscle fiber number is not exclusively determined genetically as had previously been presumed. The ability to manipulate fetal muscle fiber number by environmental factors may be possible. Proliferation and differentiation of myoblasts carries on in postnatal life; however, we were concerned with the period of gestation when the majority of differentiation takes place leading to the formation of the majority of muscle fibers. Once these fibers are formed, their numbers cannot be altered. The remaining undifferentiated myoblasts (which probably give rise to the satellite cells postnatally) are still able to proliferate and differentiate, but they fuse with existing fibers and are unable to form new fibers. The histology sections and the gene expression data suggest that the majority of fibers are formed around d 85. We therefore postulate that increasing or decreasing myoblast proliferation before this time may alter the number of fibers formed (and possibly the number of satellite cells), whereas effects after this time will not affect the numbers of fibers, but they might affect their size (hypertrophy), which depends on myoblast or satellite cell proliferation and differentiation with existing fibers.
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Implications
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This study identified the period during pregnancy when muscle cell differentiation and major fiber formation takes place in the ovine fetus. The period before differentiation and fiber formation (i.e., during proliferation) is potentially sensitive to external factors; therefore, muscle development could potentially be altered during this time period by environmental factors such as a change in nutrient supply. The results from this experiment will be used to define the periods during gestation when the muscle development of the unborn lamb may be affected by nutrient restriction of the pregnant ewe.
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Footnotes
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1 The authors thank L. Stubbins LSSC for her help with the design of the ewe diet and J. Craigon for his statistical advice. 
2 A. Fahey was funded by a Biotechnology and Biological Sciences Research Council (BBSRC) studentship.
3 Correspondence—phone: 0115 9516137; fax: 0115 951612; e-mail: Peter.Buttery{at}nottingham.ac.uk.
Received for publication December 1, 2004.
Accepted for publication May 31, 2005.
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Literature Cited
|
|---|
Applied Biosystems. 1997. User Bulletin #2: ABI PRISM 7700 Sequence Detection System, Applied Biosystems.
Bass, J. J., M. Sharma, J. Oldham, and R. Kambadur. 2000. Muscle growth and genetic regulation. Pages 227–236 in Ruminant Physiology: Digestion, Metabolism, Growth and Reproduction. CAB Int., Oxfordshire, U.K.
Brameld, J. M., A. Mostyn, J. Dandrea, T. J. Stephenson, J. M. Dawson, P. J. Buttery, and M. E. Symonds. 2000. Maternal nutrition alters the expression of insulin-like growth factors in fetal sheep liver and skeletal muscle. J. Endocrinol. 167:429–437.[Abstract]
Brameld, J. M., H. Smail, N. Imram, N. Millard, and P. J. Buttery. 1999. Changes in expression of IGF-I, IGF-II and GH receptor (GHR) mRNA during differentiation of cultured muscle cells derived from adult and fetal sheep. S. Afr. J. Anim. Sci. 29(ISRP):307–310.
Chomczynski, P., and N. Sacchi. 1987. Single step method of RNA isolation by acid guandinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 163:156–159.
Coolican, S. A., D. S. Samuel, D. Z. Ewton, F. J. McWade, and J. R. Florini. 1997. Themitogenic and myogenic actions of insulin-like growth factors utilize distinct signaling pathways. J. Biol. Chem. 272:6653–6662.[Abstract/Free Full Text]
Dauncey, M. J., and R. S. Gilmour. 1996. Regulatory factors in the control of muscle development. Proc. Nutr. Soc. 55:543–559.[Medline]
De Chiara, T. M., E. J. Robertson, and A. Efstratiadis. 1991. Parental imprinting of the mouse insulin-like growth factor II gene. Cell 64:849–859.[Medline]
Dickson, M. C., J. C. Saunders, and R. S. Gilmour. 1991. The ovine IGF-I gene-characterisation, expression and identification of a putative promoter. J. Mol. Endocrinol. 6:17–31.[Abstract]
Florini, J. R., and D. Z. Ewton. 1990. Highly specific inhibition of IGF-I stimulated differentiation by an antisense oligonucleotide to myogenin mRNA. J. Biol. Chem. 265:13435–13437.[Abstract/Free Full Text]
Florini, J. R., D. Z. Ewton, S. L. Falen, and J. J. VanWyk. 1986. Biphasic concentration dependency of stimulation of myoblast differentiation by somatomedins. Am. J. Physiol. 250:C771–C778.
Florini, J. R., D. Z. Ewton, and S. L. Roof. 1991a. Insulin-like growth factor-1 stimulates terminal myogenic differentiation by induction of myogenin gene expression. Mol. Endocrinol. 5:718–724.[Abstract]
Florini, J. R., K. A. Magri, D. Z. Ewton, P. L. James, K. Grindstaff, and P. S. Rotwein. 1991b. Spontaneous differentiation of skeletal myoblasts is dependent upon autocrine secretion of insulin-like growth factor-II. J. Biol. Chem. 266:15917–15923.[Abstract/Free Full Text]
Gerrard, D. E., C. S. Okamura, M. A. M. Ranalletta, and A. L. Grant. 1997. Developmental expression and location of IGF-I and IGF-II mRNA and protein in skeletal muscle. J. Anim. Sci. 76:1004–1011.
Harper, J. M. M., J. B. Soar, and P. J. Buttery. 1987. Changes in protein metabolism of ovine primary muscle cultures on treatment with growth hormone, insulin, insulin-like growth factor I or epidermal growth factor. J. Endocrinol. 112:87–96.[Abstract]
Hasty, P., A. Bradley, J. A. Morris, D. G. Edmondson, J. M. Venuti, E. N. Olsen, and W. H. Klein. 1993. Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene. Nature 364:501–506.[Medline]
Hynes M. A., J. J. Van Wyk, P. J. Brooks, A. J. D’Ercole, M. Jansen, and P. K. Lund. 1987. Growth hormone dependence of somatomedin-C/insulin like growth factor-I and insulin like growth factor-II messenger ribonucleic acids. Mol. Endocrinol. 1:223–242.
Isaksson, O. G., S. Eden, and J. Jansson. 1985. Mode of action of pituitary growth hormone on target cells. Annu. Rev. Physiol. 47:483–499.[Medline]
Ji, S. Q., R. L. Losinski, S. G. Cornelius, G. R. Frank, G. M. Willis, D. E. Gerrard, F. F. S. Depreux, and M. E. Spurlock. 1998. Myostatin expression in porcine tissues: Tissue specificity and developmental and postnatal regulation. Am. J. Physiol. Reg. Integ. Comp. Physiol. 44:R1265–R1273.
Joulia, D., H. Bernardi, V. Garandel, F. Rabenoelina, B. Vernus, and G. Cabello. 2003. Mechanisms involved in the inhibition of myoblast proliferation and differentiation by myostatin. Exp. Cell. Res. 286:263–275.[Medline]
Klempt, M., B. Bingham, B. H. Breier, W. R. Baumbach, and P. D. Gluckman, 1993. Tissue distribution and ontogeny of growth hormone receptor messenger ribonucleic acid and ligand binding to hepatic tissue in the mid gestation sheep fetus. Endocrinology 132:1071–1077.[Abstract]
Kambadur, R., M. Sharma, and T. P. Smith. 1997. Mutations in myostatin (GDF8) in double-muscled Belgian Blue and Pied-montese cattle. Genome Res. 7:910–916.[Abstract/Free Full Text]
Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685.[Medline]
Lee, S. J., and A. C. McPherron. 1999. Myostatin and the control of skeletal muscle mass. Curr. Opin. Gen. Dev. 9:604–607.[Medline]
Liu, J. P., J. Baker, A. S. Perkins, E. J. Robertson, and A. Efstratiadia. 1993. Mice carrying null mutations of the genes encoding insulin-like growth factor I (IGF-1) and type 1 IGF receptor (IGF1r). Cell 75:59–72.[Medline]
Magri, K. A., M. R. Benedict, D. Z. Ewton, and J. R. Florini. 1994. Negative feedback regulation of insulin-like growth factor-II gene expression in differentiating myoblasts in vitro. Endocrinology 135:53–62.[Abstract]
Maltin, C. A., M. I. Delday, K. D. Sinclair, J. Steven, and A. A. Sneddon. 2001. Impact of manipulations of myogenesis in utero on the performance of adult skeletal muscle. Reproduction 122:359–374.[Abstract]
McPherron, A. C., A. M. Lawler, and S. J. Lee. 1997. Regulation of skeletal muscle mass in mice by a new TGF-ß superfamily member. Nature 387:83–90.[Medline]
McPherron, A. C., and S. J. Lee. 1997. Double muscling in cattle due to mutations in the myostatin gene. Proc. Nat. Acad. Sci. USA 94:12457–12461.[Abstract/Free Full Text]
Musaro, A., and N. Rosenthal. 1999. Maturation of the myogenic program is induced by postmitotic expression of insulin-like growth factor I. Mol. Cell. Biol. 19:3115–3124.[Abstract/Free Full Text]
Oldham, J. M., J. A. Martyn, M. Sharma, F. Jeanplong, R. Kambadur, and J. Bass. 2001. Molecular expression of myostatin and MyoD is greater in double-muscled RT than normal-muscled cattle fetuses. Am J. Phys. Reg. Integr. Comp. Physiol. 280:R1488–R1493.
Rehfeldt, C., N. C. Stickland, I. Fiedler, and J. Wegner. 1999. Environmental and Genetic Factors as sources of variation in skeletal muscle fibre number. Basic Appl. Myo. 9:235–253.
Rosenthal, S. M., E. J. Brown, A. Brunetti, and I. D. Goldfine. 1991. Fibroblast growth factor inhibits insulin-like growth factor II (IGF-II) gene expression and increases IGF-I receptor abundance in BC3H-I cells. Mol. Endocrinol. 5:678–684.[Abstract]
Sakuma, K., K. Watanabe, M. Sano, I. Uramoto, and T. Totsuka. 2000. Differential adaptation of growth and differentiation factor 8/myostatin, fibroblast growth factor 6 and leukaemia inhibitory factor in overloaded, regenerating and denervated rat muscles. Biochim. Biophys. Acta 1497:77–88.[Medline]
Smith, J. A., A. M. Lewis, P. Wiener, and J. L. Williams. 2000. Genetic variation in the bovine myostatin gene in UK beef cattle: Allele frequencies and haplotype analysis in the south devon. Anim. Gen. 31:306–309.
Smythe, G. M., M. J. Davis, D. Paulin, and M. D. Grounds. 2001. Absence of desmin slightly prolongs myoblast proliferation and delays fusion in vivo in regenerating grafts of skeletal muscle. Cell Tissue Res. 304:287–294.[Medline]
Soumillion, A., J. H. F. Erkens, J. A. Lenstra, G. Rettenberger, and M. F. Te pas. 1997. Genetic variation in the porcine myogenin gene locus. Mamm. Genome 8:564–568.[Medline]
Tseng, B. S., S. T. Cavin, E. P. Hoffman, S. T. Iannaccone, P. Mancias F. W. Booth, and I. J. Butler. 1999. Human bHLH transcription factor gene myogenin (MYOG): Genomic sequence and negative mutation analysis in patients with severe congenital myopathies. Genomics 57:419–423.[Medline]
Yamanouchi, K., C. Soeta, N. Naito, and H. Tojo. 2000. Expression of myostatin gene in regenerating skeletal muscle of the rat and its localization. Biochem. Biophys. Res. Commun. 270:510–516.[Medline]
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Abstract
Introduction
