J. Anim Sci. 2007. 85:1247-1256. doi:10.2527/jas.2006-511
© 2007 American Society of Animal Science
Contractile protein content reflects myosin heavy-chain isoform gene expression1
A. M. Gunawan,
S. K. Park,
J. M. Pleitner,
L. Feliciano,
A. L. Grant and
D. E. Gerrard2
Department of Animal Sciences, Purdue University, West Lafayette, IN 47907
 |
Abstract
|
|---|
Muscle fiber types are classified based on contractile speed and type of metabolism. Fast-contracting fibers involve mainly glycolytic-based metabolism, whereas slow-contracting fibers involve a more oxidative type of energy metabolism. The relationship between expression of the genes controlling these functional characteristics and their relative protein abundance in porcine muscle is unknown. The objective of this study was to determine the expression of adult myosin heavy-chain (MyHC) genes and their corresponding protein content in various porcine muscles. Moreover, changes in expression of 2 genes involved in energy metabolism (glycogen synthase and citrate synthase) were determined on muscles varying in MyHC. Using real-time PCR, the relative transcript abundance was determined for the adult MyHC isoforms (types I, IIA, IIX, and IIB), glycogen synthase, and citrate synthase in the masseter (MAS), diaphragm, longissimus, cutaneous trunci, and red and white semitendinosus muscles of 7 pigs. Each muscle was subjected to SDS-PAGE analyses to determine the relative abundance of each MyHC. The relative transcript abundance of type IIB MyHC was greatest (P < 0.05) in the longissimus, white semitendinosus, and cutaneous trunci muscles, whereas type I MyHC expression was greatest (P < 0.05) in the MAS, diaphragm, and red semitendinosus muscles. Glycogen synthase gene expression was least in the MAS (P < 0.01) but exhibited a pattern similar to MyHC IIB expression across muscles. Citrate synthase transcript abundance, however, varied (P < 0.05) independently of MyHC gene expression. Expression of types I and IIB MyHC was correlated with their tissue protein content (R2 = 0.76 and 0.78, respectively), whereas type IIA and X MyHC expression did not correlate with the SDS-PAGE-determined protein content. These data show differences in MyHC gene expression across various porcine muscles and suggest that expression of these genes is reflective of the type of myosin contained within the muscle. Moreover, these data show that expression of energy-specific genes differs greatly across porcine muscles with different functions.
Key Words: myosin heavy chain • porcine • skeletal muscle
|
INTRODUCTION
|
|---|
Skeletal muscle consists of a set of fiber types differing in their ability to contract and metabolize energy. One determinant of contractile speed is the type of myosin heavy chain (MyHC) within the muscle fiber, which corresponds to fiber types classified histochemically as I, IIA, IIX, or IIB fibers (Lefaucheur et al., 1998
). 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.
|
MATERIALS AND METHODS
|
|---|
Animals
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.
Actomyosin ATPase Staining
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.
|
RESULTS
|
|---|
Figures 1 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.
View larger version (151K):
[in this window]
[in a new window]
|
Figure 1. Histochemical staining of serial cross-sections of masseter (top row), red semitendinosus (middle row), and diaphragm (bottom row) porcine skeletal muscles. Adenosine triphosphatase staining (left column) of type I, IIA, and IIX/IIB fibers resulted in dark, light, and intermediate staining, respectively. Reduced ß-nicotinamide adenine dinucleotide staining (right column) of type I, IIA, and IIX/IIB fibers resulted in dark, light, and undetectable staining, respectively.
|
|
View larger version (149K):
[in this window]
[in a new window]
|
Figure 2. Histochemical staining of serial cross-sections of longissimus (top row), white semitendinosus (middle row), and cutaneous trunci (bottom row) porcine skeletal muscles. Adenosine triphosphatase staining (left column) of type I, IIA, and IIX/IIB fibers resulted in dark, light, and intermediate staining, respectively. Reduced ß-nicotinamide adenine dinucleotide staining (right column) of type I, IIA, and IIX/IIB fibers resulted in dark, light, and undetectable staining, respectively.
|
|
View larger version (49K):
[in this window]
[in a new window]
|
Figure 3. Representative SDS-PAGE analysis (A) of masseter (MAS), red semitendinosus (RST), diaphragm (DIA), longissimus (LM), white semitendinosus (WST), and cutaneous trunci (CT) porcine skeletal muscles of 7 pigs. Bottom, middle, and top bands contained type I, IIB, and IIX/A myosin heavy chain (MyHC), respectively. Relative abundance (B) of type I, IIA/X, and IIB MyHC protein in the MAS, RST, DIA, LM, WST, and CT porcine skeletal muscles. Means bearing different letters differ (P < 0.05).
|
|
Control ß-actin gene expression did not vary across muscles (data not shown) validating our qPCR techniques. However, various MyHC isoforms varied (P < 0.05) with the muscle type. Relative transcript abundance of type I MyHC was 54-fold greater in RST than in WST (Figure 4A). 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.
View larger version (17K):
[in this window]
[in a new window]
|
Figure 4. Relative expression (Ct, threshold cycle) of type I (A), IIA (B), IIX (C), and IIB (D) myosin heavy-chain (MyHC) genes in porcine masseter (MAS), red semitendinosus (RST), diaphragm (DIA), longissimus (LM), white semitendinosus (WST), and cutaneous trunci (CT) muscles (n = 7). Means bearing different letters differ (P < 0.05).
|
|
Expression of the CS gene was greatest (P < 0.01) in DIA and did not differ from RST, WST, and CT, but was 2-fold lower in LM and MAS (Figure 5A). 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).
View larger version (13K):
[in this window]
[in a new window]
|
Figure 5. Relative citrate synthase (A) and glycogen synthase (B) gene expression (Ct, threshold cycle) in porcine masseter (MAS), red semitendinosus (RST), diaphragm (DIA), longissimus (LM), white semitendinosus (WST), and cutaneous trunci (CT) muscles (n = 7). Means bearing different letters differ (P < 0.05).
|
|
Figures 6A and 6B 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).
View larger version (18K):
[in this window]
[in a new window]
|
Figure 6. Relationship between the relative protein abundance and gene expression of type I (A), IIB (B), and IIA/X (C) myosin heavy-chain (MyHC) isoforms in porcine masseter, red and white semitendinosus, diaphragm, longissimus, and cutaneous trunci muscles. Each data point in panels A and B represents the mean of 7 pigs, whereas each data point in panel C represents a sample from 1 pig.
|
|
|
DISCUSSION
|
|---|
Adult porcine skeletal muscle consists of 4 different fiber types (I, IIA, IIX, IIB; Lefaucheur et al., 1998), 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.
|
Footnotes
|
|---|
1 Purdue University Agricultural Research Programs Journal Paper No. E2006-511. 
2 Corresponding author: dgerrard{at}purdue.edu
Received for publication July 28, 2006.
Accepted for publication November 8, 2006.
|
LITERATURE CITED
|
|---|
Bass, A., D. Brdiczka, P. Eyer, S. Hofer, and D. Pette. 1969. Metabolic differentiation of distinct muscle types at the level of enzymatic organization. Eur. J. Biochem. 10:198–206.[Medline]
Beermann, D. H., V. K. Fishell, K. Roneker, R. D. Boyd, G. Armbruster, and L. Souza. 1990. Dose-response relationships between porcine somatotropin, muscle composition, muscle fiber characteristics, and pork quality. J. Anim. Sci. 68:2690–2697.[Abstract]
Brooke, M. H., and K. K. Kaiser. 1970. Three "myosin adenosine triphosphatase" systems: The nature of their pH lability and sulfhydryl dependence. J. Histochem. Cytochem. 18:670–672.[Medline]
Chaudhary, R., B. P. Chowdhary, I. Harbitz, I. Gustavsson, and C. T. Evans. 1992. Localization of the citrate synthase (CS) gene to the p12–p13 bands of chromosome 5 in pigs by in situ hybridization. Hereditas 117:39–43.[Medline]
Chomczynski, P., and N. Sacchi. 1987. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162:156–159.[Medline]
Cox, R. D., and M. E. Buckingham. 1992. Actin and myosin genes are transcriptionally regulated during mouse skeletal muscle development. Dev. Biol. 149:228–234.[CrossRef][Medline]
da Costa, N., R. Blackley, H. Alzuherri, and K. C. Chang. 2002. Quantifying the temporospatial expression of postnatal porcine skeletal myosin heavy chain genes. J. Histochem. Cytochem. 50:353–364.[Abstract/Free Full Text]
Depreux, F. F. S., A. L. Grant, and D. E. Gerrard. 2002. Influence of halothane genotype and body-weight on myosin heavy chain composition in pig muscle as related to meat quality. Livest. Prod. Sci. 73:265–273.[CrossRef]
Depreux, F. F. S., C. S. Okamura, D. R. Swartz, A. L. Grant, A. M. Brandsetter, and D. E. Gerrard. 2000. Quantification of myosin heavy chain isoform in porcine muscle using an enzyme-linked immunosorbent assay. Meat Sci. 56:261–269.[CrossRef]
Guth, L., and F. J. Samaha. 1970. Procedure for the histochemical demonstration of actomyosin ATPase. Exp. Neurol. 28:365–367.[CrossRef][Medline]
Karlsson, A., A. C. Enfalt, B. Essen-Gustavsson, K. Lundstrom, L. Rydmer, and S. Stern. 1993. Muscle histochemical and biochemical properties in relation to meat quality during selection for increased lean tissue growth rate in pigs. J. Anim. Sci. 71:930–938.[Abstract]
Lefaucheur, L., and D. E. Gerrard. 2000. Muscle fiber plasticity in farm mammals. Proc. Am. Soc. Anim. Sci. http://www.asas.org/jas/symposia/proceedings/0307.pdf Accessed Nov. 8, 2006.
Lefaucheur, L., R. K. Hoffman, D. E. Gerrard, C. S. Okamura, N. Rubinstein, and A. Kelly. 1998. Evidence for three adult fast myosin heavy chain isoforms in type II skeletal muscle fibers in pigs. J. Anim. Sci. 76:1584–1593.[Abstract/Free Full Text]
Lefaucheur, L., D. Milan, P. Ecolan, and C. Le Callennec. 2004. Myosin heavy chain composition of different skeletal muscles in Large White and Meishan pigs. J. Anim. Sci. 82:1931–1940.[Abstract/Free Full Text]
Lefaucheur, L., E. Patrick, L. Plantard, and N. Gueguen. 2002. New insights into muscle fiber types in the pig. J. Histochem. Cytochem. 50:719–730.[Abstract/Free Full Text]
Mandarino, L. J., R. L. Printz, K. A. Cusi, P. Kinchington, R. M. O’Doherty, H. Osawa, D. K. Granner, and R. A. DeFronzo. 1995. Regulation of hexokinase II and glycogen synthase mRNA, protein, and activity in human muscle. Am. J. Physiol. 269:E701–E708.[Medline]
McKoy, G., M.-E. Leger, F. Bacou, and G. Goldspink. 1998. Differential expression of myosin heavy chain mRNA and protein isoforms in four functionally diverse rabbit skeletal muscles during pre- and postnatal development. Dev. Dyn. 211:193–203.[CrossRef][Medline]
Ono, Y., M. B. Solomon, C. M. Evock-Clover, N. C. Steele, and K. Maruyama. 1995. Effects of porcine somatotropin administration on porcine muscles located within different regions of the body. J. Anim. Sci. 73:2282–2288.[Abstract]
Peter, J. B., R. J. Barnard, V. R. Edgerton, C. A. Gillespie, and K. E. Stempel. 1972. Metabolic profiles of three fiber types of skeletal muscle in guinea pigs and rabbits. Biochemistry 11:2627–2633.[CrossRef][Medline]
Pette, D., and R. S. Staron. 2000. Myosin isoforms, muscle fiber types, and transitions. Microsc. Res. Tech. 50:500–509.[CrossRef][Medline]
Picard, B., J. Leger, and J. Robelin. 1994. Quantitative determination of type I myosin heavy chain in bovine muscle with anti myosin monoclonal antibodies. Meat Sci. 36:333–343.[CrossRef]
Schiaffino, S., and C. Reggiani. 1996. Molecular diversity of myofibrillar proteins: Gene regulation and functional significance. Physiol. Rev. 76:371–423.[Abstract/Free Full Text]
Schiaffino, S., and C. Reggiani. 1994. Myosin isoforms in mammalian skeletal muscle. J. Appl. Physiol. 77:493–501.[Abstract/Free Full Text]
Sepponen, K., N. Koho, E. Puolanne, M. Ruusunen, and A. R. Poso. 2003. Distribution of monocarboxylate transporter isoforms MCT1, MCT2 and MCT4 in porcince muscles. Acta Physiol. Scand. 177:79–86.[CrossRef][Medline]
Talmadge, R. J., and R. R. Roy. 1993. Electrophoretic separation of rat skeletal muscle myosin heavy-chain isoforms. J. Appl. Physiol. 75:2337–2340.[Abstract/Free Full Text]
Talmadge, R. J., R. R. Roy, and V. R. Edgerton. 1993. Muscle fiber types and function. Curr. Opin. Rheumatol. 5:695–705.[Medline]
Weiss, A., and L. A. Leinwand. 1996. The mammalian myosin heavy chain gene family. Annu. Rev. Cell Dev. Biol. 12:417–439.[CrossRef][Medline]
Top
Abstract
INTRODUCTION
