Muscle atrophy is a physiological response to diverse physiological and pathological conditions that trigger muscle deterioration through specific cellular mechanisms. Despite different signals, the biochemical changes in atrophying muscle share many common cascades. Muscle deterioration as a physiological response to the energetic demands of fish vitellogenesis represents a unique model for studying the mechanisms of muscle degradation in non-mammalian animals. A salmonid microarray, containing 16,006 cDNAs, was used to study the transcriptome response to atrophy of fast-switch muscles from gravid rainbow trout compared with sterile fish. Eighty-two unique transcripts were upregulated and 120 transcripts were downregulated in atrophying muscles. Transcripts having gene ontology identifiers were grouped according to their functions. Muscle deterioration was associated with elevated expression of genes involved in the catheptic and collagenase proteolytic pathways; the aerobic production, buffering, and utilization of ATP; and growth arrest; whereas atrophying muscle showed downregulation of genes encoding a serine proteinase inhibitor, enzymes of anaerobic respiration, muscle proteins as well as genes required for RNA and protein biosynthesis/processing. Therefore, gene transcription of the trout muscle atrophy changed in a manner similar to mammalian muscle atrophy. These changes result in an arrest of normal cell growth, protein degradation, and decreased glycolytic cellular respiration that is characteristic of the fast-switch muscle. For the first time, other changes/mechanisms unique to fish were discussed including genes associated with muscle atrophy.
- Oncorhynchus mykiss
several physiological and pathological conditions can cause muscle atrophy by triggering unique responses through distinct cellular stimuli. Transcriptional mechanisms of muscle wastage are well characterized at the transcriptome level in mammalian models as a response to nutritional restriction (7), fasting, severe diseases, including cancer, renal failure, diabetes (41, 42), and sepsis (42), as well as muscle disuse or denervation (4, 57). Some studies have dealt with fish muscle atrophy at the level of individual genes/mechanisms (46, 51, 58, 60–62). The molecular basis of mammalian muscle atrophy has received considerable attention in the literature (25, 30). Losing muscle mass results from elevated rate of proteolysis and decreased rate of protein synthesis. Despite the significant amount of literature (30, 32, 42), the specific roles of the proteolytic systems in degrading mammalian muscle still are unclear. Moreover, we recently showed that fish use different muscle degrading proteolytic strategies compared with mammals (58). The ubiquitin-proteasome pathway, which is responsible for the bulk of proteolysis in mammalian muscle atrophy (31, 42), is not involved in the vitellogenesis-induced atrophy (58) and is even downregulated (46) in fasting-induced fish muscle degradation. Several external signals that regulate muscle proteolysis have been identified, but little is known about intramuscular signal transduction mechanisms that couple external stimuli to the regulation of protein degradation (32).
The primary objective of this study was to characterize global gene expression profiles in atrophying muscle of rainbow trout (RBT). The vitellogenesis-induced muscle atrophy represents a unique nonmammalian muscle degradation model, because fish are ectothermic animals that swiftly use proteins as oxidative substrates (69) and have lower metabolic rate than endothermic animals. Muscle wastage in RBT is a response to a dominant physiological process rather than a pathological condition of dying fish seen in semelparous fish species, such as salmon (Oncorhynchus nerka and Oncorhynchus keta). this study, we used this model 1) to determine whether a common transcriptional program controls muscle degradation in mammalian and nonmammalian animals and 2) to distinguish any unique piscine mechanisms. This study represents the first high-throughput analysis of expression changes that address muscle degradation for a nonmammalian model. Characterizing a common transcriptional profile in organisms with different genetic backgrounds, environmental and metabolic adaptations should increase our understanding of the decisive and evolutionary conserved mechanisms associated with muscle atrophy. In addition, these analyses identify general and specific features of muscle catabolism among different species and name targets of genetic manipulations that favoring muscle growth in aquaculture species.1
MATERIALS AND METHODS
Fish and Muscle Sampling
Mature fertile (diploid) and sterile (triploid) female RBT, Oncorhynchus mykiss (∼500 g), were collected from Flowing Springs Trout Farm (Delray, WV) during the spawning season in early October. Fish were cultured in identical raceways receiving water from a common spring at 13 ± 3°C. Fish were fed ad libitum (Zeiglar Gold; Zeigler Bros., Gardeners, PA) via demand feeders. No difference of food consumption was noticed between groups. Fish had access to feed when sampled. As confirmed by dissection, fertile fish were gravid with a gonado-somatic index (GSI = ovary weight/fish weight × 100) of 15.8 ± 0.3 (n = 5). The GSI of sterile fish was 0.3 ± 0.2 (n = 5). White muscle samples (∼20 g) from five fish of each group were collected from the dorsal musculature and flash frozen in liquid nitrogen and stored at −80°C until RNA extraction. Following tissue removal, fish were eviscerated, and ribs and vertebral column were removed in a filleting procedure. The filleted muscle and sample weights were combined and expressed as a percentage of the whole fish weight as an indicator of the muscle mass. A portion of the muscle was used for proximate composition and texture analyses.
Muscle Texture and Composition Analyses
Muscle shear force was measured as described previously (58, 59). Shear force was measured with a 5-blade Kramer shear attachment using a Texture Analyzer (model TA-HDi; Texture Technologies, Scarsdale, NY), and results were expressed as g force/g of sample. Muscle proximate composition (moisture, protein, fat) was determined according to Association of Official Analytical Chemists-approved procedures (3).
Total RNA was isolated from each fish (5 fish/group) using the TRIzol reagent (Invitrogen, Carlsbad, CA) according to the manufacturer's instruction. Concentrations of isolated RNA were determined by measuring absorbance at 260 nm. The integrity of RNA was determined by agarose gel electrophoresis. Poly(A) mRNA was purified using Oligotex mRNA Mini Kit (Qiagen, Valencia, CA) according to the manufacturer's instruction.
Microarrays, cDNA Labeling, and Hybridization
A salmonid microarray containing cDNAs representing 16,006 genes selected from Atlantic salmon and RBT expressed sequence tag databases (70) was used to profile changes in gene expression of atrophying muscles. Microarray chips were obtained from Drs. Ben Koop and William Davidson at the University of Victoria through the Genomic Research on Atlantic Salmon Project (GRASP). A compete list of the genes on the array is available at http://web.uvic.ca/cbr/grasp/. The microarray has been validated as a useful tool for RBT studies (70). Gene expression levels were determined by comparing the amount of mRNA transcript present in the experimental sample (fertile fish) to the control (sterile fish). RNAs isolated from each experimental fish were run on separate microarrays in independent experiments, with no pooling. A total of 10 fish were used in the microarray experiments (5 replicates × 2 groups). Fluorophors (Cy3 and Cy5) were randomly assigned to RNA from each of the atrophying and nonatrophying muscles to limit the dye effect. cDNA labeling and microarray hybridization procedures were essentially as described (75). In brief, 0.8 μg of mRNA from each trout muscle were used as a template in reverse transcription reactions incorporating amino-allyl dUTP into the cDNA using oligo-d (T) primer and Superscript II reverse transcriptase (Invitrogen). The synthesized cDNAs from atrophying (fertile fish) and nonatrophying muscles (sterile fish) were differentially labeled using N-hydroxysuccinate-derived Cy3 or Cy5 dyes (GE Healthcare, Piscataway, NJ). Labeled cDNAs were purified using a PCR purification kit (Qiagen) to remove unincorporated dyes. The Cy3- and Cy5-labeled cDNAs were then combined and concentrated down to 20 μl using a Microcon-30 concentrator (Millipore, Billerica, MA) followed by addition of 130 μl of Slidehyb 3 solution (Ambion, Austin, TX). Microarray hybridizations were performed on a Tecan HS400 automated microarray hybridization station (Tecan US, Durham, NC). The slides were placed on the machine at 60°C for 2 min followed by prehybridization at 55°C for 30 min with prehybridization solution (5× SSC, 1% SDS, 1% BSA) under medium agitation. After a brief washing at 60°C for 1 min, differentially labeled cDNAs in hybridization buffer (∼145 μl) were injected into the hybridization chamber. The hybridizations were carried out for 3 h at 60°C followed by another 13 h at 55°C. Arrays were washed twice in 2× SSC, 01% SDS, followed by twice in 0.1× SSC, 0.1% SDS at room temperature. Following two more washes in 0.1× SSC, the slides were rinsed in water and dried by centrifugation.
Microarray Scanning and Data Analysis
ScanArray Lite microarray scanner was used to scan arrays and ScanArray Express software (Perkin Elmer, Wellesley, MA) was used to process array images, align spots, integrate robot-spotting files with the microarray images, and quantify spots. Preprocessing and normalization of data were accomplished using the R-project statistical environment (http://www.r-project.org) and Bioconductor (http://www.bioconductor.org) through the GenePix AutoProcessor (GPAP) website (http://darwin.biochem.okstate.edu/gpap). Data were preprocessed by: 1) removing data points where signal intensities in both channels was less than a baseline threshold value of 200, 2) calculating and subtracting local background fluorescence values from all feature intensities, 3) Log2-transforming the background subtracted Cy3/Cy5 ratios, 4) averaging the technical replicates within and across the replicates, and 5) defining spots that are larger or smaller than 2 SDs from the mean as outliers and eliminating them from calculation of the final average Log2 ratios within and across array replicates. All hybridizations were also subjected to manual review to ensure flagging and exclusion of all unacceptable spots. Following preprocessing, the expression results were normalized using global LOWESS normalization. For each spot, B-statistic, t-statistic, P value (probability), and M value [log2 (cy5/cy3)] were calculated. Spots with twofold change or more (represented by an M value ≤ −1 or ≥ +1) were considered differentially expressed using P value < 0.05. Five experiments were conducted. Microarray data were deposited (according to Microarray Gene Expression Data Society Standards) in the NCBI Gene Expression Omnibus (GEO, http://www.ncbi.nlm.nih.gov/geo/) with the series accession number (GSE4787).
Quantitative Real-time PCR Analysis
Quantitative real-time PCR was used to confirm the expression of nine differentially expressed genes identified by microarray experiments. Total RNA, isolated from muscle samples (n = 5/group), using TRIzol reagent, was further purified using an RNA clean-up kit according to the manufacturer's protocol (Zymo Research, Orange, CA). We converted 2 μg of each RNA sample to cDNA using Superscript II reverse transcriptase (Invitrogen). To ensure RNAs were free of genomic DNA, negative control cDNAs were prepared by reverse transcription reactions without adding the reverse transcriptase. Real-time PCR primers were designed based on each gene sequence (Table 1) using Primer3 software (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi). Quantitative PCR was performed in duplicate for each cDNA sample on a Bio-Rad iCycler iQ Real-Time PCR Detection System using iQ SYBR Green Supermix (Bio-Rad, Hercules, CA) in 25-μl reaction volumes containing 300 nM of each primer and cDNA derived from 0.2 μg of total RNA. The stable probe RBT β-actin gene in the microarray was chosen as an endogenous control for normalization of the real-time PCR analysis. Standard curves for each gene and the endogenous control were constructed using 10-fold serial dilutions of the corresponding plasmid. Standard curves were run on the same plate with the samples. Threshold lines were adjusted to intersect amplification lines in the linear portion of the amplification curve and cycles to threshold (Ct) were recorded. For each sample, the amount of target gene and endogenous reference was determined from the appropriate standard curve. The amount of the target gene was divided by the amount of reference gene to obtain a normalized target value. Mean differences in expression levels were reported as relative fold changes. This was done by designating the control group (nonatrophying muscles) as a calibrator and dividing the mean of treatment group (atrophying muscles) by the mean of the calibrator (calibrator mean divided by itself equals one) (60). Mean differences in gene expression levels were determined by t-test using a statistical analysis package, SigmaStat version 3.11 (Aspire Software International, Leesburg, VA).
Muscle Atrophy as a Response to Vitellogenesis
Atrophying muscle of fertile fish had 11% less extractable muscle (g/body wt) and 11% less protein content compared with nonatrophying muscle of sterile fish (Fig. 1, A and B; P < 0.01). The decrease in total protein content was associated with a 4% increase in water content (P < 0.01). Muscle fat content was not affected (P > 0.05). Shear force of the nonatrophying muscle of sterile fish was greater (P < 0.05) than the atrophying muscle of the fertile fish (data not shown). We did not measure muscle glycogen content. However, assessment of the postmortem muscle pH revealed that nonatrophying muscle had a lower (P < 0.01) ultimate pH (6.41 ± 0.04) than atrophying muscle (6.61 ± 0.03). These data suggest that atrophic muscle may contain less glycogen as the substrate for anaerobic metabolism thereby resulting in less lactic acid accumulation and higher ultimate pH.
Identification of differentially expressed genes associated with muscle atrophy using a 16k cDNA microarray defined 82 upregulated and 120 downregulated unique transcripts in atrophying muscles. The majority of the genes (∼99%) were not differentially expressed (P > 0.05, M value > −1 or < 1). Most of the genes lie within the ± 2-fold change range. Gene ontology classifications of the differentially expressed genes gave functional gene sets that represent a specific program of transcriptional changes. Expression of nine genes identified by microarray as differentially expressed in atrophic muscle was confirmed by quantitative real-time PCR analysis. These genes were selected according to their biological functions. Each gene represents one functional gene set as described below. All nine genes showed statistically significant changes (P < 0.05) in atrophying muscle vs. nonatrophying muscles (Fig. 2).
Genes Involved in Protein Metabolism
Expression of cathepsin L, D, and S was upregulated in atrophying muscle (Table 2). In addition, a gene encoding the serine proteinase inhibitor, serpin was downregulated in atrophying muscle. Surprisingly, expression of the multicatalytic proteasome system that is responsible for the bulk of mammalian proteolysis was not altered. Transcription of 34 genes encoding subunits of the proteasome pathway was not affected. In addition, the 26S proteasome subunit-β type 5 and the regulatory subunit 4 were downregulated. Moreover, the 26S proteasome, non-ATPase regulatory subunits 3, 12, and 14 were downregulated in atrophying muscle (Table 2). The lack of a significant role for the proteasome pathway in RBT protein degradation was supported by less mRNA of the ubiquitining genes: ubiquitin-activating enzyme E1c and ubiquitin-conjugating enzymes E2, E2N, and E2L3 in atrophying muscles. Expression of the collagenase-3 enzyme was upregulated in atrophying muscle (Table 2). Dipeptidyl-peptidase I and an aminopeptidase were also upregulated in atrophying RBT muscle.
Myofibrillar and muscle regulatory protein.
In atrophying RBT, muscle mRNAs of myosin heavy chain, and α-actin were markedly depleted. In addition, mRNA abundance of the myosin alkali light chain 4 was downregulated in atrophying muscle (Table 3). Similarly, expression of the muscle regulatory protein troponin I and tropomyosin-α 1 and 5 was downregulated (Table 3). The unconventional myosin type I α-gene was downregulated as well. Expression of other important cytoskeletal intrasarcomeric proteins like α-actinin, myomesin (skelemin), and H-protein was not affected. Expression of the cytoskeletal intersarcomeric proteins, such as desmin, talin, ankyrin, and β-spectrin, was not affected. Similarly, expression of the muscle regulatory proteins tropomodulin and the trimeric protein members, troponins C, and T was not affected.
Extracellular matrix/cell adhesion proteins.
Atrophying muscle had reduced expression of genes encoding important extracellular matrix and connective tissue proteins: collagen types I α1, 2, 3, and x; SPARC (osteonectin/BM-40), an important basement membrane protein; and fasciclin-like adhesion proteins ig-h3, a collagen-binding protein (Table 3).
With the loss of muscle mass, expression of the cytoskeletal proteins tubulin-α 1, 2, 4, and 6 and tubulin-β was downregulated (Table 3). In addition, keratin type II 6A, a cytoskeletal protein, and prefoldin-4, a chaperone that facilitates folding of tubulin and actins (45), exhibited reduced expression in atrophying muscle. The list of the downregulated genes also includes an intracellular hyaluronan-binding protein, a member of a recently identified proteins involved in proliferation of smooth muscle cells and fibroblasts (15), and a gastrulation-specific G12-like gene, a microtubule-stabilizing protein (5).
RNA Processing, Protein Biosynthesis, and Modification
Degenerating RBT muscle had a reduced abundance of transcripts involved in transcription and RNA processing, protein biosynthesis, posttranslational modification, and intracellular protein translocation (Table 4). The RNA processing genes include transcription initiation factors, mRNA splicing, and polyadenylation factors. The protein biosynthesis list comprises ribosomal assembly proteins, translation initiation and elongation factors, and heat shock proteins chaperonins, in addition to nucleoplasms that play important role in packaging of rRNA and ribosomal proteins during ribosome formation. Surprisingly, an ATP-dependent, RNA helicase WM6, a member of the DEAD-box family that is induced in mammalian muscle wastage, was downregulated, while another transcript of RNA helicase-related protein was upregulated in atrophying RBT muscle. In addition, calnexin, which is a component of the ER chaperone system of glycoproteins, was downregulated. Atrophying muscle showed declined expression of genes encoding rate-limiting steps in protein folding and posttranslational modification. Two of those genes, the peptidyl-prolyl cis-trans isomerase A and B (cyclophilins), are ubiquitous enzymes that catalyze the cis-trans isomerization of peptidylprolyl bonds. The downregulated posttranslational modification genes include protein disulfide-isomerase A3 and protein-lysine 6-oxidase, as well. Accumulation of transcripts of the intracellular protein transport process was also impaired in degenerating muscle, including: 1) β- and γ-subunits of the Sec61 protein, which is an endoplasmic reticulum (ER) cotranslational protein transport system (65); 2) Sar1 protein, which promotes vesicle budding from the endoplasmic reticulum (39); 3) AP-1 adaptor binding protein, which has been identified on clathrin-coated vesicles (54); 4) multidrug resistance-associated protein 1, which transports a wide variety of xeno- and endobiotics (9); and 5) glutamine synthetase, which is consistently highly expressed in mammalian muscle degradation.
Genes Involved in Glucose Metabolism
RBT atrophying muscle had decreased transcription of genes encoding many of the glycolytic enzymes, indicating reduction in glucose utilization (Table 5). Transcripts for enzymes involved in the early and intermediate steps of glycolysis had decreased expression in atrophying muscle. These enzymes include glucose-6-phosphate isomerase, fructose-bisphosphate aldolase A, triosephosphate isomerase, glyceraldehyde-3-phosphate dehydrogenase, and phosphoglycerate kinase 1. Similarly, mRNAs of enzymes catalyzing late steps in glycolysis, α- and β-enolase were reduced. Furthermore, mRNAs of the lactate dehydrogenase, an enzyme that acts on products of glycolysis, was also diminished (Table 5). Other members of the glycolytic pathway, including the highly regulated phosphofructokinase, were not affected. Noteworthy, expression of glyceraldehyde-3-phosphate dehydrogenase, a gene commonly used as a housekeeping gene, with expected stable expression, was significantly decreased in atrophying muscle.
Tricarboxylic acid cycle, oxidative phosphorylation, and hexose monophosphate shunt.
Atrophying muscle of RBT had increased mRNAs of the tricarboxylic acid (TCA) enzymes isocitrate dehydrogenase and succinate dehydrogenase (Table 5). In addition, many genes involved in the oxidative phosphorylation process were differentially expressed in atrophying muscle. Several genes belonging to complexes of the electron transport pathway: complex I (NADH-ubiquinone oxidoreductases), complex III (ubiquinol-cytochrome c oxidoreductases), complex IV (cytochrome c oxidase), and complex V ATP synthase, showed differential expression. There were more upregulated genes of the oxidative phosphorylation process than downregulated ones. Transcript accumulation of the hexose monophosphate shunt enzyme glucose-6-phosphate 1-dehydrogenase (G6PD) mRNA was higher in the atrophying RBT muscle (Table 5).
ATP buffering and utilization.
Expression of the creatine kinases, M-type, B-type, and the mitochondrial acidic type, were consistently upregulated (Table 5). Expression of the sarcoplasmic/endoplasmic reticulum calcium ATPase 2 was upregulated.
Transcription Factors/Signal Transduction Regulators
Many genes involved in diverse array of signal transduction cascades that regulate cell cycle progression and apoptosis were upregulated in atrophying muscle (Table 6), including: 1) the growth arrest and DNA-damage-inducible proteins (GADD)45β and GADD45γ genes; 2) Sestrin 1 (PA26), which is a p53-regulated protein; 3) protein kinase Cδ, which is believed to be a proapoptotic gene; 4) death-associated protein (DAP)-1, which is involved in mediating TNF-induced cell death; 5) TOB protein, which is an antiproliferative cell cycle regulator (47); 6) RalBP1, which binds Ral proteins [Ral, with its upstream molecule Ras, is a GTPase involved in regulation of myogenic cell migration (66) and inhibits myogenesis (36)]; 7) Ras GTPase-activating-like protein IQGAP1, a multifunctional protein plays a key part in regulating E-cadherin-mediated cell adhesion (56); 8) CCAAT/enhancer binding protein delta (C/EBPδ), activated in mammalian skeletal muscle damage (12) and by a glucocorticoid-dependent mechanism in septic muscle (55); 9) muscle-specific RING finger protein 3 (MuRF3); and 10) γ-aminobutyric acid receptor associated protein, which is upregulated in mammalian muscle wastage (7, 41).
On the other hand, RBT degenerating muscle downregulated the expression of many genes (Table 6) involved in several signal transduction cascades that regulate cell cycle progression, including: 1) the anaphase-promoting subunit 1, ubiquitin-protein ligase, and 2) thyroid hormone receptor interactor 3.
Genes of Miscellaneous and Unknown Functions
Selenoproteins genes are the major biological form of selenium in animals. Oxidoreductases are the only selenoproteins with know function, involved in multiple metabolic pathways. Two glutathione peroxidases showed elevated expression in atrophying muscle, indicating possible roles in free radical scavenging and/or maintenance of intracellular redox status (Table 7). Selenoproteins S and M showed reduced expression in atrophying muscle. On the other hand, selenoprotein P expression was elevated, indicating possible isoform shifting. Transposons activated in RBT microarray studies by external stimuli, such as toxicity and stress, and correlated with a group of genes implicated in defense response, signal transduction, and regulation of transcription (37). Two transposases had elevated expression in atrophying RBT muscle (Table 7). Serotransferrin, which is responsible for the iron transport was also upregulated in atrophying muscle (Table 7). In addition, 64 transcripts of unknown annotation showed differential expression in atrophying muscle (Table 8). These are candidates for further investigation, looking for key genes in muscle degradation.
Muscle Atrophy in Response to Vitellogenesis
Our results indicate that RBT during vitellogenesis lose a significant proportion of the muscle mass and mobilize a substantial amount of muscle proteins likely to accumulate large amounts of yolk in developing ova (51). The reduction in muscle protein is coupled with an increase in muscle water content and muscle softness, suggesting that the broken-down proteins are replenished with water (58). Similar patterns of muscle wastage have been described in other fish, such as sockeye salmon, Oncorhynchus nerka (50), and chum salmon, Oncorhynchus keta (51, 74). These changes are consistent with a general pattern of muscle wastage, which supports using RBT as an ideal model to elucidate the molecular mechanisms of muscle catabolism in fish. Transcriptional profiling, using a 16k cDNA microarray, defined a group of 202 of the genes differentially expressed in RBT vitellogenesis-induced muscle atrophy. This number represents ∼1% of the total number of probes present on the chip. Using gene ontology to classify these genes, based on molecular function, resulted in gene sets that represent a specific program of transcriptional changes that are largely similar to the well-studied programs of mammalian muscle atrophy (41, 57). In addition, a number of new and unexpected genes, perhaps unique to the RBT wastage process, are reported here for the first time.
Genes Involved in Protein Metabolism
Expression of cathepsin L, D, and S were upregulated in atrophying muscle. This was coupled with downregulation of the serine proteinase inhibitor, serpin. This synchronized expression confirms our previous observation that cathepsins, especially cathepsin L, constitute the major protein catabolic pathway in vitellogenesis-associated muscle degradation (58). Cathepsin L was reported among a common set of genes, termed atrogins, that were induced in four catabolic states of mammalian muscles (41). Consequently, cathepsin L represents a well evolutionarily conserved protease associated with muscle atrophy.
On the other hand, expression of 34 genes encoding subunits of the proteasome pathway did not change in response to the vitellogenesis-induced muscle degeneration. In addition, the 26S proteasome subunit-β type 5 and regulatory subunit 4 were downregulated. Moreover, the 26S proteasome non-ATPase regulatory subunits 3, 12, and 14 were downregulated in atrophying muscle. The unregulated expression of the proteasome pathway genes in RBT protein degradation, leading to our suggestion that the proteasome pathway is not important to RBT muscle wastage, is further supported by the reduced expression of the ubiquitination genes; ubiquitin-activating enzyme E1c, ubiquitin-conjugating enzyme E2, E2N, and E2L3. The multicatalytic proteasome system is responsible for the bulk of mammalian proteolysis (31, 42), and several proteasome genes were denoted as atrogins, genes associated with muscle catabolic states (41). These results confirm previous reports that RBT proteasome pathway, unlike in mammals, has no significant role in piscine protein turnover (46, 58). Together these results indicate altered mechanisms of the muscle protein degradation in fish compared with mammals, with cathepsins being the dominant proteases in mobilizing fish muscle proteins. Transcripts of four calpain genes were not changed, likely indicating a limited role of the calpains in vitellogenesis-induced muscle degeneration. Previously, we showed that calpains are involved in proteolysis of proteins to fuel metabolism in RBT during starvation in muscle (61) and liver (63). In addition, we recently reported that mRNA of calpastatin, the calpain inhibitor, is associated with muscle growth in RBT (62). Consequently, our current results may imply that vitellogenesis stimulates muscle proteolysis through signaling transduction cascades that are different from those used by starving RBT.
In mammals, the muscle atrophy-associated genes (41) included very few muscle-specific proteins and neither of the major contractile proteins, myosin nor actin. Similarly, denervated rat muscle had downregulated expression of only muscle regulatory proteins (4). Our results reveal reduced transcription of the major muscle contractile proteins, myosin and actin, which constitute ∼60% and ∼20% of muscle proteins, respectively. In addition, myosin alkali light chain 4, troponin I, tropomyosin-α 1 and 5, and the unconventional myosin type I α-gene were downregulated. Our results are consistent with similar studies showing that both soluble and structural proteins were degraded in spawning-induced salmon muscle degeneration (50). Van den Thillart (69) reported that fish utilize proteins as oxidative substrates. This indicates that fish, unlike mammals, during substantial negative nitrogen balance may sacrifice the important contractile proteins myosin and actin, using them as proteins reservoir. It may also explain how fish, unlike mammals, can tolerate long periods of starvation.
Many of the extracellular matrix proteins that play critical roles in the dynamic structural changes during development and tissue remodeling, as well as in maintenance of mechanical properties and homeostasis of the muscle, had downregulated expression in atrophying muscle. Extracellular matrix and connective tissue genes including collagens, SPARC and fasciclin-like adhesion proteins, ig-h3, had reduced expression in the atrophic RBT muscle. Collagen is the most abundant protein found in animals, accounting for 20–30% of total protein (18). The downregulated expression of major muscle proteins collagens myosin and actin may point to a substantial negative nitrogen balance resulting from the energy demand by developing ova. Extracellular matrix acts as a structural scaffold for alignment of myofibrils. Diminution of the extracellular matrix, indicated by downregulation of collagen and upregulation of collagenase 3 enzyme, would elicit the reduction in shear force of the atrophying muscle and, in turn, deterioration of muscle food quality. SPARC (osteonectin/BM-40) is an important component of the basement membrane proteins, a specialized extracellular matrix that supports cell adhesion, migration, and proliferation. SPARC binds calcium and collagens, and it can modulate cell-matrix interactions, thereby altering cell shape, growth, and differentiation (17). Our routine histological examination showed that atrophying RBT muscle had irregular shapes, widening of intermyofibrillar spaces, and less cell-cell adhesion than nonatrophying muscle (data not shown). Downregulation of SPARC and collagens may explain, in part, these structural changes. Similarly, fasciclin-like adhesion proteins ig-h3 and β-ig-h3 are collagen-binding proteins and play important roles in collagen interactions in various tissues (23). The β-ig-h3 gene has been induced in several cell lines by TGF-β1 (23), indicating possible involvement in mediating some signals of this multifunctional growth modulator. Reduced expression of the ig-h3 proteins suggests there may be limited ability of the atrophying muscle cells to respond to growth signals.
Degenerating muscle exhibited downregulation in expression of genes encoding the important cytoskeletal proteins. Altered transcription of the cytoskeletal proteins including tubulin is not a common feature of the mammalian muscle degradation program described by Lecker and coworkers (41). Batt and colleagues (4) noted that tubulin is downregulated only after prolonged denervation characterized by irreversible muscle impairment, profound atrophy, myocyte death, and fibrosis, whereas short-term denervation, associated with fully reversible changes, did not lead to tubulin alteration. These results suggest that altered tubulin expression is associated only with severe muscle degeneration.
RNA Processing, Protein Biosynthesis, and Modification
Degenerating RBT muscle suppressed the overall protein synthesis process. The reduced expression of genes involved in protein biosynthesis occurred at multiple levels, including RNA transcription and processing, translation, posttranslational modification, and intracellular protein translocation. The RNA processing genes include transcription initiation factors and splicing and polyadenylation factors. The translation list includes ribosomal assembly proteins, translation initiation and elongation factors, heat shock proteins and chaperonins and nucleoplasms that play important roles in packaging of rRNA and ribosomal proteins during ribosome formation. An ATP-dependent RNA helicase, WM6 protein, a member of the DEAD-box family, was also downregulated. Similar helicases are important in the ribosomal subunits biogenesis (14). In contrast to our finding, the WM6 helicase is constantly induced in catabolic states of mammalian muscles (41), although another RNA helicase-related protein was upregulated in atrophying muscle. This discrepancy suggests altered mechanisms of ribosomal biosynthesis in mammals vs. fish. The atrophying muscles showed declined expression of genes encoding rate-limiting steps in protein folding and posttranslational modification including the peptidyl-prolyl cis-trans isomerases A and B (cyclophilins), which are enzymes catalyzing the cis-trans isomerization of peptidylprolyl bonds (21). The posttranslational list also includes protein disulfide-isomerase A3 and protein-lysine 6-oxidase. The latter is an extracellular enzyme that catalyzes the cross-linking of collagen and elastin through oxidative deamination of lysine or hydroxylysine side chains (68). Weakening of the cross-linking of these proteins may further explain the decreased muscle integrity and quality of RBT because of the vitellogenesis effect. The posttranslational list also includes calnexin, which is a component of the ER chaperone system that ensures the proper folding of newly synthesized glycoproteins (72). Transcripts of the intracellular protein transport process were also impaired in degenerating muscle. As anticipated, glutamine synthetase was induced in degenerating muscle. Glutamine synthetase increases dramatically in atrophic muscle as a result of chronic exposure to glucocorticoids (26) or denervation (16). Glutamine synthetase catalyzes de novo glutamine synthesis from glutamate and ammonia, thus maintaining glutamine homeostasis under conditions of increased glutamine demand by other tissues (13) or as a result of its limited supply. Muscle atrophy induces glutamine efflux, thereby depleting muscle glutamine stores (28).
Genes Involved in Glucose Metabolism
The gene expression pattern of the degenerating muscle of RBT revealed a decrease in expression of seven glycolytic enzymes in addition to the lactate dehydrogenase enzyme that act on products of glycolysis. Decreased expression of glycolytic enzymes in atrophying muscle may be due to insufficient glycogen as a substrate of anaerobic metabolism. Postmortem muscle pH was lower ultimately in nonatrophied than in atrophied muscle. Less glycogen in atrophic muscle may result in less lactic acid accumulation and higher ultimate pH. Glucose sparing in RBT may result from insulin resistance because of the prolonged energetic challenge from the reproductive demand. Reduced transcription of genes involved in glucose utilization is a common feature of muscle atrophy induced by fasting, the systemic diseases diabetes, cancer cachexia, and renal failure (41) and muscle disuse (57). Insulin resistance is a common feature of all forms of muscle degradation and is the direct trigger of accelerated muscle proteolysis (32, 41). Glucocorticoids alone can induce muscle atrophy (24) and seem to be essential for atrophies caused by fasting, renal failure, and diabetes (41). Mommsen (51) suggested that the spawning-induced salmon muscle degenerative mechanisms involve glucocorticoids. Fish spawning (11, 38), and intense exercise (49) increase activity of the pituitary-interrenal axis, increasing cortisol availability. This observation suggests that altered gene expression of glycolytic pathway enzymes may be subsequent to insulin resistance caused by prolonged partial anorexia or as a downstream effect of elevated levels of adrenal glucocorticoids.
Atrophying RBT muscle had increased mRNAs for the TCA cycle enzymes isocitrate dehydrogenase and succinate dehydrogenase. Several genes, belonging to different complexes of the electron transport pathway, were differentially expressed with some being upregulated and others downregulated, probably reflecting specific remodeling in atrophying muscle. There were more electron transport genes upregulated than downregulated. In addition, subunit 6 of the mitochondrial ATPase, an enzyme complex that catalyzes coupling of proton flux (oxidative phosphorylation) to ATP synthesis (52), had elevated expression in atrophying RBT muscle. Furthermore, expression of several isoforms of creatine kinases was consistently upregulated in atrophying muscle. Creatine kinases act as a ATP buffer catalyzing the transfer of phosphate between ATP and various phosphogens (e.g., creatine phosphate) and thereby play a central role in energy transduction in tissues with large and fluctuating energy demands like skeletal muscle. Likewise, expression of the sarcoplasmic/endoplasmic reticulum calcium ATPase 2 was upregulated. This enzyme catalyzes ATP hydrolysis coupled with the translocation of calcium from the cytosol to the sarcoplasmic reticulum, thereby facilitating regulation of muscle contraction/relaxation. Together these changes point to unanticipated enhanced competence for aerobic ATP production, buffering, and utilization. In disagreement with our findings, expression of genes encoding proteins of mitochondrial energy production was reduced in catabolic states of mammalian muscle degradation (41, 57). The list of these genes includes TCA enzymes, NADH oxidoreductase, ATP synthases, and mitochondrial creatine kinases. This contradiction suggests that changes of the aerobic pathway are not conserved between terrestrial and aquatic animals and may be not general features of atrophying muscles. Fish rapidly use proteins as oxidative substrates (69), and salmonids encountering prolonged hypoxic conditions may sustain reduced reproductive success (33). Consequently, the proposed elevation of aerobic metabolism in RBT atrophic muscle may be to cope with the elevated gluconeogenesis indicated by the increased rate of proteolysis.
Accumulation of glucose-6-phosphate 1-dehydrogenase (G6PD) mRNA was higher in atrophying RBT muscle. G6PD plays a key role in the production of ribose 5-phosphate and the generation of NADPH in the hexose monophosphate shunt, bypassing glycolysis. The products of the hexose monophosphate shunt are the reduced form of NADP, which may supply lipid biosynthesis to support the developing ova, and the C5 sugars, which may contribute to biosynthesis of nucleotides and key muscle proteins. The net chemical effect of this pathway is the same as the combined operation of glycolysis and citric acid cycle. The shunt pathway is not equally important in all animal tissues. It is more significant in aerobic tissues and unimportant in skeletal muscle where the main reaction sequence is glycolysis. The altered expression of the genes involved in glucose metabolism suggests shifting toward aerobic cellular respiration of the atrophying muscle. Metabolic changes in gene expression appear to be coordinated in the direction of a fast-to-slow fiber type transformation. However, quantitative expression analysis of the slow-specific myosin heavy chain gene indicated no transformation of muscle fiber types (data not shown). Oxidative capacity increased in sarcopenia associated with low food intake, and proportions of the slow oxidative fibers increased at the expense of fast glycolytic fibers (71). Consequently, changes in atrophying RBT muscle may not imply increased incorporation of aerobic respiration proteins into muscle but, rather, to relative preferential loss of anaerobic fibers.
Transcription Factors/Signal Transduction Regulators
RBT degenerating muscle revealed upregulated expression of many genes involved in a diverse array of signaling transduction cascades that regulate cell cycle progression and apoptosis. These genes are 1) GADD45β and GADD45γ genes, which encode for nuclear proteins associated with cell growth control, apoptotic cell death, and the cellular response to DNA damage and muscle denervation (1, 2, 57) and whose expressions are transactivated by p53 (67); 2) sestrin 1 (PA26), which is similar to the muscle atrogin PA26-T2 (41) and is p53-regulated as well; and 3) protein kinase Cδ, whose activation has a critical proapoptotic role in cardiac responses following ischemia (53) and is working downstream of the p53 response to the DNA damage-induced apoptosis (27). Elevated expression of p53 in diabetes (20) and taurine deficiency (19) is associated with cardiomyopathy. Expression of the important fish muscle degrading enzyme, cathepsin D, is likely controlled by the tumor suppressor gene p53 (73). Coordinated expressions of the GADD45s, sestrin 1, protein kinase Cδ, and cathepsin D genes suggest that an intramuscular signal transduction cascade, including the p53 gene, is modulating RBT muscle catabolism. Expression of the p53 gene was not responsive to muscle atrophy in RBT or mammals consistent with a translational or posttranslational mechanism for the p53 role (44). Caiozzo and coworkers (8) reported that increased expression in the key cell cycle inhibitor, GADD45, in denervated muscle reflects an inhibition of satellite cell proliferation and/or a special form of apoptosis that results in loss of myonuclei known to occur under atrophic conditions. The Caiozzo et al. report is in agreement with our previous results that apoptotic caspase enzymes are upregulated (58) and with our current observation that number of myonuclei decreases (histological observation, data not shown) in RBT atrophying muscles. RBT degenerating muscle had downregulated expression of the cell cycle regulator anaphase-promoting subunit 1 gene, which is a member of a multisubunit ubiquitin-protein ligase that binds the mitotic regulators securin and cyclins substrates and promotes their destruction (10). Synchronized expression of the above-mentioned cell cycle regulators suggests that RBT muscle atrophy is, at least in part, a growth suppression program.
In addition to growth suppression, RBT atrophic muscle exhibited signs of enhanced apoptosis. This was exemplified by upregulation of DAP-1, which is involved in mediating TNF-induced cell death (43). There is a significant body of literature on the role of TNF-α in muscle proteolysis (32). TNF-α is an important cytokine trigger of muscle wasting associated with disease states, but not disuse atrophy. It seems that the TNF-α pathway is activated during RBT atrophy, working either parallel or upstream of the proposed p53/GADD45 signaling.
Atrophic muscle had elevated expression of MuRF3. MuRF3 is a myogenic regulator of the muscle microtubule network and is required for myogenesis (64). MuRF1 and MuRF2 are of significant interest because of their role in signal transduction and transcriptional regulation during muscle atrophy. MuRF1 is a RING finger containing ligase reported as an important atrogin (41) required for muscle atrophy programs (32). Deletion of MuRF1 in mice inhibits skeletal muscle atrophy (6). MuRF1 is able to bind the transcription factor glucocorticoid modulatory element binding protein-1 (48), hence modulating transcription of the glucocorticoid-responsive genes. The role of the MuRF atrogin family seems to be evolutionarily conserved and consistent with an essential function in modulating muscle degeneration. MuRF proteins can be used as biomarkers of muscle atrophy. MuRF1 is a ubiquitin ligase that degrades cardiac troponin-1 in a proteasome-dependent manner (34). Our results indicate that the proteasome proteolytic pathway is not activated in RBT vitellogenesis-induced atrophy (58). Further detailed experiments are needed to characterize how MuRF3 may induce RBT muscle atrophy.
RBT degenerating muscle had downregulated expression of thyroid hormone receptor interactor 3, which is a member of the steroid hormone receptor superfamily and can activate HNF-4α, a transcription factor. It plays an important role in regulating the expression of genes involved in a variety of metabolic pathways in pancreatic β-cells and perhaps glucose metabolism (29). Thyroid hormone receptor induced myocardial growth by a mechanism involving interaction with MAP kinases (35). Low food intake is associated with reduced plasma thyroid hormone and fewer muscle thyroid hormone receptors (22). These results point to a potential role for thyroid hormone in mediating some catabolic responses of the vitellogenesis-associated RBT muscle degradation.
To conclude, an adaptive molecular profile of the atrophying RBT skeletal muscle can be generally sketched into four clear trends.
The first trend is upregulated protein turnover of the myofibrillar, extracellular, and cytoskeleton proteins. The process involves synchronized transcriptional regulation. Transcripts of the myosins, α-actin, tropomyosins, troponin I, collagens, osteonectin, tubulins, and prefoldin are downregulated. However, transcripts of the enzymes degrading these proteins, such as cathepsins, collagenase, and peptidases, were upregulated. Consistent with this, the serine proteinase inhibitor serpin is downregulated. The changes described in this trend generally fits the changes denoted in the mammalian models. Nevertheless, expression of the proteasome proteolytic pathway, which is responsible for the mammalian bulk proteolysis and contains several muscle atrogins, was not affected in our study. These results confirm the previous reports that ruled out the significance of the proteasome pathway in piscine protein turnover (46, 58).
The second trend is suppression of the overall protein synthesis process. This trend involves all steps of protein biosynthesis from transcription and RNA processing to translation, posttranslational modification, and intracellular protein translocation. The genes clustered under this category generally outline a similar trend as observed in mammalian muscle atrophy models. The increase in protein turnover induces gluconeogenesis. Consistent with that, glutamine synthetase was induced in atrophying muscle suggesting a potential usage as a molecular biomarker for testing muscle degradation. Atrophic muscle must manage oxidative stress and cellular breakdown; regulated gene expression consistent with this need was the observed upregulation of the glutathione peroxidases. Glutathione peroxidase has been shown to be involved with maintaining membrane integrity (40) as well as free radical scavenging and/or maintenance of intracellular redox status.
The third trend is decrease in the expression of the glycolytic enzymes. This trend represents a common characteristic of all forms of muscle atrophy, and it is believed to be due to the insulin resistance. We propose that a prolonged caloric challenge or increased corticosteroids because of the vitellogenesis stresses may result in a case of insulin resistance in RBT. On the other hand, our results point to an unanticipated enhanced competence for aerobic ATP production, buffering, and utilization. These changes are contrary to a general trend of suppressing the mitochondrial energy production reported in catabolic states of mammalian muscle degradation. This contradiction suggests that changes in the aerobic pathway are not a general feature of atrophying muscles.
Finally, a well-regulated expression pattern of diverse arrays of signal transduction cascades that regulate cell cycle progression and apoptosis was noted. We propose that an intramuscular signal cascade modulating RBT muscle catabolism is centralized around the p53 gene. Consequently, RBT muscle atrophy is, at least partly, regulated through a growth suppression program perhaps coupled with apoptosis. Any growth arrest/apoptotic changes described here should be seen as reversible changes. After spawning, RBT will regain lost muscle by releasing the growth arrest mechanisms and reversing the protein synthesis/degradation ratio. These regeneration mechanisms are currently under investigation in our lab. Despite of the lack of involvement of the proteasome pathway in RBT muscle degeneration, expression of the ubiquitin ligase atrogin MuRF3 was upregulated. The evolutionary conserved MuRF atrogin family may has an essential function in modulating muscle degeneration and may be used as universal biomarkers of muscle atrophy. Several genes with differential expression patterns that seem unique to the fish model were reported for the first time. Further characterization of these genes may reveal new insights into the regulation of muscle growth and protein turnover in aquaculture species.
This investigation was supported by USDA Cooperative Agreement no. 58-1930-5-537 and Hatch Project no. 427. The paper is published with the approval of the Director of the West Virginia Agricultural and Forestry Experiment Station as scientific paper no. 2964.
↵1 The 2nd International Symposium on Animal Functional Genomics was held May 16–19, 2006 at Michigan State University in East Lansing, Michigan and was organized by Jeanne Burton of Michigan State University and Guilherme J.M. Rosa of University of Wisconsin-Madison (see meeting report by Drs. Burton and Rosa, Physiol Genomics 28: 1-4, 2006.).
Address for reprint requests and other correspondence: J. Yao, Div. of Animal and Veterinary Sciences, West Virginia Univ., Morgantown, WV 26506-6108(e-mail:).
Article published online before print. See web site for date of publication (http://physiolgenomics.physiology.org).
- Copyright © 2007 the American Physiological Society