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Department of Animal Sciences, Purdue University, West Lafayette, IN 47907
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Abstract |
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 Top Abstract INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: myosin heavy chain • porcine • skeletal muscle
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INTRODUCTION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Each MyHC isoform is expressed by a member of a large, multigene family (Weiss and Leinwand, 1996). Fibers containing mostly type I MyHC are the slowest contracting fibers, whereas type IIB MyHC-containing fibers contract with the greatest velocity.
Muscle fibers are "plastic," meaning they can change their function to accommodate myriad environmental cues. In nontransitory muscle fibers, MyHC transcript abundance reflects contractile protein abundance (Lefaucheur et al., 2002). These data support the concept that the MyHC composition in adult pig muscle is transcriptionally controlled, similar to that shown for other species (Cox and Buckingham, 1992; Schiaffino and Reggiani, 1996). Little information exists regarding the relationship between muscle MyHC content and relative gene expression in the pig.
Furthermore, muscle type is classified according to the type of energy metabolism used (glycolytic or oxidative). Fast-contracting fibers possess predominately glycolytic metabolism, whereas slow-contracting fibers use mainly oxidative metabolism, but the relationship between genes controlling energy metabolism and contractile speed is not known. This is of interest to those wishing to optimize pork production, because there is a strong positive relationship between the number of fast-contracting glycolytic fibers and lean growth rate (Karlsson et al., 1993; Depreux et al., 2002), yet this attribute may be negatively associated with pork quality (reviewed by Lefaucheur and Gerrard, 2000). Therefore, the overall objective of this study was to determine the fiber type-specific gene expression in several porcine muscles.
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MATERIALS AND METHODS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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The Purdue Animal Care and Use Committee approved all procedures for the care and use of pigs. Seven pigs (115 ± 5 kg) obtained from the Purdue University Research and Education Center were slaughtered and processed according to normal industry procedures. Masseter (MAS), diaphragm (DIA), longissimus, cutaneous trunci (CT), and red (RST) and white (WST) semitendinosus muscle samples were taken within 10 min postexsanguination. Samples for real-time PCR were frozen in liquid nitrogen and stored at –80°C. Samples for histochemistry were frozen in 2-methybutane cooled in liquid nitrogen and then stored at –80°C.
Total RNA Preparation
Total RNA was isolated from porcine skeletal muscle using the single-step method (Chomczynski and Sacchi, 1987). Briefly, 100 mg of skeletal muscle powdered in liquid nitrogen was placed into 3 mL of a denaturing solution containing 4 M guanidium isothiocyanate (Sigma, St. Louis, MO), 25 mM sodium citrate (pH 7), 0.5% sarcosyl, and 0.1 M ß-mercaptoethanol, in a 50-mL conical tube held on ice. Tissue was homogenized using a Polytron homogenizer (Brinkman Instruments, New York, NY) at speed 6, then transferred to a fresh 15-mL conical tube. Next, 300 µL of 2 M sodium acetate (pH 4) was added and the tubes were vortexed. Protein and lipids in the preparation were extracted by sequentially adding 3 mL of phenol saturated with diethyl pyrocarbonate-treated water (pH 4.3) and 600 µL of chloroform-isoamyl alcohol (40:1, vol/vol). After thorough vortexing, the tubes were placed on ice for 15 min. Samples were centrifuged at 10,000 x g at 4°C for 20 min to separate the aqueous and organic phases. The aqueous phase (upper) was carefully transferred to a fresh 15-mL conical tube. Attention was taken to avoid disturbing the interphase. Precipitation of RNA was facilitated by adding 3 mL of isopropanol and holding the samples at –20°C for 1 h. Precipitated RNA was then collected by centrifugation at 10,000 x g at 4°C for 30 min. The pellets were then redissolved in 0.9 mL of denaturing solution. Next, 0.9 mL of isopropanol was added, and the RNA was allowed to precipitate at –20°C for 1 h. Precipitated RNA was then collected by centrifugation at 21,000 x g at 4°C for 30 min. Isopropanol was discarded, and the pellets were washed twice with 1 mL of 75% ethanol (vol/vol). The pellets were air dried for 10 min and resuspended in 50 µL of TE-8 [1 M Tris-HCl (pH 8) and 0.2 M EDTA (pH 8) in diethyl pyrocarbonate-treated water].
RNA Quantification
Total RNA isolated from skeletal muscle was measured using the Ribogreen quantification kit (Molecular Probes, Eugene, OR). The RNA was first treated with 4 U of recombinant DNase (Ambion, Indianapolis, IN) in 2 µL of a 10x DNase buffer containing 100 mM Tris-HCl (pH 7.5), 25 mM MgCl2, 5 mM CaCl2, and 3 µL of nuclease-free water (Ambion), and digested for 30 min at 37°C. Five microliters of DNase inactivation reagent (Ambion), a resin that binds DNase, was added to the reaction and mixed by flicking the tubes. Ribonucleic acid was then loaded into 0.22-µm Spin-X centrifuge tube filters (VWR International, West Chester, PA) and centrifuged at 10,000 x g for 1 min.
A 1-µL sample of purified RNA was diluted with 249 µL of TE-8 containing 200 mM Tris-HCl and 20 mM EDTA (pH 7.5; Ambion). Next, 75 µL of TE buffer and 100 µL of Ribogreen reagent were combined in each well of a 96-well microplate. Twenty-five microliters of each prepared RNA sample was added to the reagents on the microplate. A GENios Pro fluorometer (Tecan, Durham, NC) was used to measure the fluorescent signal at 480 (excitation) and 520 (emission) nm.
cDNA Synthesis
Ribonucleic acid was next reverse transcribed to cDNA. First, cDNA master mix was prepared from 5 U of Moloney murine leukemia virus (M-MLV) reverse transcriptase (Invitrogen, Carlsbad, CA), 0.5 U of SU-PERase-In (Ambion), 10 mM DTT (Invitrogen), and 5x first-strand buffer (Invitrogen). Total RNA (0.5 µg) in 5 µL of nuclease-free water was added to 100 ng of random hexamers/µL and 100 µM of each dNTP. Samples were denatured at 65°C for 5 min and then chilled to 4°C on ice. Four microliters of cDNA master mix was added to the denatured RNA and sequentially incubated at 25°C for 10 min, 37°C for 50 min, 70°C for 10 min, and chilled to 4°C on ice. Finally, cDNA samples were diluted with nuclease-free water so that gene quantification was based on 50 ng of cDNA per 5 µL.
Real-Time PCR
Relative transcript abundance was determined using quantitative real-time PCR (qPCR). First, gene-specific master mix was prepared from 2x iQ SYBR Green Supermix containing 100 mM KCl, 40 mM Tris-HCl (pH 8.4), 0.4 mM of each dNTP, 50 units of iTaq DNA polymerase/mL, 6 mM MgCl2, SYBR Green I 7.5 µL, 20 nM fluorescein stabilizers (BioRad Laboratories, Hercules, CA), 10 pmol/µL of each gene-specific forward and reverse primer, and nuclease-free water.
Briefly, 5-µL samples of cDNA were added in duplicate to a 96-well microplate. Next, 10 µL of the gene-specific master mix was added to each well containing cDNA. Primer sequences for the MyHC isoforms, glycogen synthase (GS), citrate synthase (CS), and ß-actin are shown in Table 1. Real-time PCR was run in 15-µL reactions for 40 cycles using an iCycler real-time PCR detection system (BioRad) according to optimized gene-specific protocols as described in Table 2. Amplified qPCR products were sequenced to validate primer specificity. The first cycle in the log-linear region of amplification in which a significant increase in fluorescence was detected above background was designated the threshold cycle (Ct). Amplification of highly expressed genes is detected at a lower cycle number than genes expressed at a lower level, which are not detected until after many cycles of PCR; therefore, the Ct value and gene expression are inversely related.
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Serial, 10-µm cross-sections of each muscle sample were made using a Microm HM 500M cryostat (Microm, Walldorf, Germany). The chamber temperature was –25°C. Serial cross-sections were used to localize actomyosin ATPase using a modification of the procedure of Guth and Samaha (1970). Briefly, muscle cross-sections were preincubated for 5 min at room temperature in a solution (pH 4.35) containing 0.1 M HCl, 70 mM sodium acetate, and 14 mM sodium barbital. Sections were initially incubated for 45 min at 37°C in a solution (pH 10.4) containing 2 mM ATP, 40 mM calcium chloride, 70 mM sodium chloride, and 60 mM sodium hydroxide. Sections were next incubated for 3 min at room temperature in 60 mM calcium chloride, 3 min at room temperature in 80 mM cobalt chloride, and 1 min at room temperature in 14 mM ammonium sulfide. Between each incubation step, the tissue sections were rinsed 3 times with distilled water. Sections were allowed to dry for 1 h, mounted, and coverslipped.
NADH Staining
Muscle cross-sections were incubated for 90 min at 37°C in a solution (pH 7.0) containing 4 mM nitro blue tetrazolium, 0.2 M Tris buffer, 50 mM magnesium chloride, and 2 mM NADH. Sections were incubated in water for 5 min at room temperature. Sections were fixed in 4% formalin for 15 min, and then rinsed with distilled water, allowed to dry, and coverslipped.
SDS-PAGE
Muscle MyHC abundance was determined by SDS-PAGE using procedures reported by Talmadge and Roy (1993). Approximately 100 mg of muscle tissue was powdered in liquid nitrogen using a mortar and pestle. The MyHC was isolated with a high-ionic-strength buffer containing 0.3 M KCl, 0.1 M KH2PO4, 40 mM EDTA, and 50% glycerol (pH 6.5). Homogenates were vigorously stirred on ice for 15 min. Samples were then centrifuged at 10,000 x g for 20 min at 4°C. Supernatants were mixed with an equal volume of glycerol. Concentration of the isolated protein was determined using a BCA protein assay kit (Pierce, Rockford, IL). Samples were suspended in 2x Laemmli buffer containing 10% SDS, 10% ß-mercaptoethanol, and 20% glycerol in 0.5 M Tris (pH 6.8), and heated in boiling water for 5 min. Extracted samples were loaded on a gel in a sample buffer containing 0.1 M Tris, 150 mM glycine, 0.1% SDS, and 0.001% ß-mercaptoethanol. Five-microgram samples of protein from each sample were resolved using SDS-PAGE. Stacking gels were composed of 30% glycerol, 4% acrylamide-N,N-methylene-bis-acrylamide (bis; 50:1), 0.5 M Tris (pH 6.8), 4 mM EDTA, and 0.4% SDS. The resolving gel consisted of 30% glycerol, 8% acrylamide-bis (50:1), 0.4 M Tris (pH 8.8), 0.1 M glycine, and 4% SDS. Gels were run on a Mini-Protein II dual slab cell electrophoretic system (BioRad) for 30 h at 70 V. Gels were stained using Coomassie blue, and band density was quantified using Kodak 1D image analysis software (Kodak, New Haven, CT).
Statistical Analysis
Data were analyzed using the GLM procedure (SAS Inst. Inc., Cary, NC) with muscle as the main effect and Ct or band intensity as the independent variable in the model. Data were reported as least squares means, and an ANOVA was used to determine differences among the means.
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RESULTS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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and 2 show representative cross-sections stained for myofibrillar ATPase (left column) and NADH (right column) for the muscles used in this study. Note the variation in the relative number of fibers differentially stained by these 2 classic fiber-typing protocols. Figure 3A shows a representative SDS-PAGE gel of the corresponding muscles used in this study. Even though we were not successful in separating all 4 adult MyHC in these muscles, type I (Figure 3A, lower band) and 2 type II (Figure 3A, upper band) MyHC bands were clearly discernible using the accepted protocol for separating MyHC in other species. Scans of these gels revealed that the MAS, RST, and DIA muscles contained greater (P < 0.05) relative amounts of type I MyHC, whereas type II MyHC was more abundant in the LM, WST, and CT muscles (Figure 3B). Although not quantified, SDS-PAGE data corroborated histochemical myofibrillar ATPase staining and showed that dark-staining, slow-twitch fibers were clearly more evident in the MAS, RST, and DIA muscles compared with the LM, WST, and CT muscles.
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). Compared with WST, the DIA, MAS, LM, and CT had 37-, 14-, 9-, and 4-fold greater (P < 0.01) type I gene expression, respectively. Type IIA MyHC, the predominant isoform contained in fast-twitch oxidative fibers, was also differentially (P < 0.01) expressed in various muscles (Figure 4B). The greatest (P < 0.001) type IIA MyHC expression was observed in the MAS and was 10-fold greater than abundance in the LM. Type IIA MyHC mRNA abundance was 6- and 4-fold greater in the RST and DIA than in the LM, respectively. Expression of the type IIX MyHC gene was least (P < 0.001) in MAS (Figure 4C) but was 150-fold greater (P < 0.0001) in the remaining muscles. Myosin heavy-chain type IIB expression was greatest (P < 0.01) in the LM and WST (Figure 4D). Similarly to MyHC type IIX, MAS expressed the least (P < 0.001) type IIB MyHC, which was 2-fold lower than LM.
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). The greatest (P < 0.05) GS gene expression was observed in the LM, yet was not expressed differently in the RST and WST. In the MAS, DIA, and CT, however, GS expression was greatly reduced compared with the LM (Figure 5B).
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show that the relative expression of types I and IIB MyHC in each muscle was highly correlated with the amount of types I and IIB protein in the skeletal muscle studied (R2 = 0.78 and 0.76, respectively). No relationships between the expression and abundance of the remaining adult MyHC (IIA and IIX) were observed across the muscles studied (Figure 6C).
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DISCUSSION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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), which are the culmination of differential expressions of specific structural proteins and metabolic enzymes (Talmadge et al., 1993; Schiaffino and Reggiani, 1996). To accommodate the ambulatory needs of the whole organism and to provide specific functions within the body, muscles must retain a plastic nature to adapt to prolonged environmental insults or various hormonal states. The heterogeneous nature of muscle fibers collectively provides each muscle with a unique range in which to function and allows the muscle the ability to "fine-tune" itself to a wide array of physiological requirements. Understanding the process of how muscles change their functional abilities in response to various environmental or management systems may lead to novel approaches to improve animal production or, in the case of human well-being, may lead to therapies to improve the quality of life. Unfortunately, defining the fiber type composition in muscle using traditional histological approaches is time-consuming and may not accurately reflect the true functional composition of the muscle. Data presented herein show that MyHC abundance, one of the primary classifiers used in assigning muscle fiber type, mirrors its gene expression for at least 2 of the 4 adult isoforms. These data suggest that MyHC genes are transcriptionally controlled, like those in other species (Cox and Buckingham, 1992; Schiaffino and Reggiani, 1994, 1996; McKoy et al., 1998), and that MyHC expression data may be a valid alternative to more traditional methodologies for predicting muscle fiber type composition.
Electrophoretic separation supports the view that fast- and slow-twitch muscles contain different amounts of type I and II MyHC. Myofibrillar ATPase and NADH staining suggested that muscle fiber types differed among the different porcine muscles studied. Specifically, the RST and MAS have a higher proportion of type I fibers and use more oxidative metabolism, whereas the WST, LM, and CT are composed of predominately type IIB fibers and possess less oxidative metabolism than their slower contracting counterparts (Picard et al., 1994; Depreux et al., 2000). Of the fast-twitch muscles studied, LM cross-sections appeared to have the greatest proportion of type IIB fibers.
Qualitative SDS-PAGE data confirmed these observations, especially regarding expression of the type I and IIB MyHC (Figure 3A and B
). The most disappointing data were the relationships, or lack thereof, between the type IIX and IIA MyHC and their respective mRNA abundance (Figure 6C). However, this relationship may simply be a function of the inability to separate these proteins using traditional SDS-PAGE protocols (Talmadge and Roy, 1993). For example, in previous work we showed that the WST contains a significant amount of immunoreactive type IIX MyHC, whereas the RST contains a large amount of type IIA MyHC (Depreux et al., 2002). Given that these isoforms are not discernible using SDS-PAGE protocols, both MyHC species yielded a significant dominant upper MyHC band (Figure 3A). Consequently, both muscles would appear to have elevated type IIX or IIA MyHC but vastly different levels of expression. Thus, we believe the lack of relationship between expression and protein content of type IIA and IIX MyHC is related to limitations in separating the 2 highly homologous proteins. Certainly, the possibility exists that the small muscle sample used to isolate RNA may have resulted in a nonhomogeneous representation of the MyHC transcript; however, this seems unlikely.
Regardless of this understandable discrepancy, our data support the idea that the deep muscles that function to maintain posture possess more type I fibers than do the more glycolytic muscles, such as the WST (Ono et al., 1995; Depreux et al., 2000). Consistent with this argument is the fact that we observed greater type I MyHC transcripts in the RST compared with the WST and CT muscles. This same trend was observed by Beermann et al. (1990) and Schiaffino and Reggiani (1994, 1996), who reported, using immunocytochemistry and traditional fiber-type staining protocols, that nearly half of the fibers in RST were type I, whereas only 4% of the fibers in the WST stained as type I muscle fibers.
Moreover, deeper muscles that act to maintain posture in a nonfatigable manner had greater type IIA MyHC expression than those superficial muscles that are normally thought to provide a quick and strong, yet fatigable, response to a stimulus. On the other hand, type IIX MyHC, a fast-twitch fiber type specific myosin isoform, was expressed at similar levels across all muscles except for the MAS, where it was present at extremely low levels. The low abundance of type IIX MyHC gene transcripts strongly suggests that few, if any, type IIX fibers exist in the MAS muscle, supporting its extremely oxidative, slow-contracting classification, at least in large ruminants (Picard et al., 1994).
Given that mispriming increases with cycle number, great care must be taken when interpreting expression data using this technology; it is possible that we overestimated expression of this gene in this and other muscles possessing relatively high Ct values. Other inconsistencies between our data and that of other fiber type-specific studies are complicated by the fact that much of the past research has relied heavily on classical fiber-typing protocols, which may have led to erroneous fiber-type composition data. In fact, detailed studies by Lefaucheur et al. (2004) showed that traditional histochemical approaches used to classify muscle fibers in porcine LM are clearly inadequate, given that there are really 4 adult MyHC isoforms rather than the 3 histochemically defined types. Relatively high levels of the fast-twitch type IIB MyHC transcripts in porcine LM and WST are consistent with the results of Lefaucheur et al. (2004) and strongly rebut the thesis that only fast-contracting myosin isoforms are present in small mammals (Pette and Staron, 2000). These data show that the bulk of MyHC gene expression in porcine muscle is reflective of MyHC protein composition, a muscle fiber characteristic predominantly used to classify the muscle fiber type composition.
In an attempt to understand the control of energy metabolism among muscles differing in fiber type composition, we determined the expression of 2 genes that may be useful in defining fast- and slow-twitch fiber populations (Bass et al., 1969; Peter et al., 1972). Glycogen synthesis is related to glucose metabolism in skeletal muscle and could therefore be an indicator of fast-contracting fibers. However, GS activity is regulated by phosphorylation events, allosteric activation, and cytosolic calcium concentrations. In fact, Mandarino et al. (1995) showed that administration of insulin had no effect on GS expression or protein content but dramatically affected enzyme activity. Thus, it was not surprising that GS gene expression only partially reflected changes in type IIB MyHC gene expression.
Type I fibers are classically known to have greater oxidative capacity than type II fibers (Brooke and Kaiser, 1970), and CS is a key regulator of aerobic energy production in cells (Chaudhary et al., 1992). Sepponen et al. (2003) found that the MAS is one of the most oxidative muscles in pigs, whereas the LM is the least oxidative. Surprisingly, CS had an expression pattern unlike any MyHC. In fact, the MAS had a relative level of CS gene expression comparable to the LM, an extremely glycolytic muscle. However, given that most enzymes are not transcriptionally regulated, further speculation about using these genes as markers for defining fiber type is unwarranted.
Data from these experiments showed that expression of the 4 adult MyHC isoforms in porcine skeletal muscles differed greatly with their anatomical location and function. Moreover, expression of these genes was highly correlated with the relative amount of the corresponding protein contained within each muscle. Given the amount of time required to conduct classical muscle fiber-typing studies and the difficulty of separating these protein isoforms in porcine muscle, MyHC gene expression may be a reliable alternative to defining muscle fiber type composition in porcine skeletal muscles. Further studies will be necessary to validate the utility of type IIA and IIX gene expression for predicting muscle MyHC composition. Use of the expression of glycolytic or oxidative enzymes, especially GS and CS, may have little utility in attaining such a goal, given that enzyme activity is independent of gene expression. Understanding how various physiological cues and management schemes change the expression of specific muscle contractile genes could lead to a better understanding of porcine skeletal muscle growth and meat quality development.
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Footnotes |
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2 Corresponding author: dgerrard{at}purdue.edu
Received for publication July 28, 2006. Accepted for publication November 8, 2006.
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LITERATURE CITED |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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