Journal of Applied Physiology AJP: Renal Physiology
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J Appl Physiol 99: 2379-2387, 2005; doi:10.1152/japplphysiol.00778.2005
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Myofiber degeneration/regeneration is induced in the cachectic ApcMin/+ mouse

Kristen A. Mehl,1,3 J. Mark Davis,1 Franklin G. Berger,2,3 and James A. Carson1,3

1Division of Applied Physiology, Exercise Science Department, 2Department of Biological Sciences, and 3Center for Colon Cancer Research, University of South Carolina, Columbia, South Carolina

Submitted 30 June 2005 ; accepted in final form 28 July 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Cachexia is characterized as an inflammatory state induced by the cancer environment, which is accompanied by the loss of muscle and fat mass. Well-investigated mechanisms of cachexia include the suppression of myofiber protein synthesis and the induction of the protein degradation. However, it is not well characterized whether chronic inflammation during cachexia induces myofiber degeneration, which contributes to muscle mass loss and decreased functional capacity. The purpose of this study was to determine whether ApcMin/+ mice, which demonstrate a chronic systemic inflammatory state due to an intestinal tumor burden, undergo cachexia and whether the myofibers exhibit signs of degeneration and/or regeneration. Six-month-old female ApcMin/+ body weight decreased 21% compared with C57BL/6 mice and was not the result of blunted growth. ApcMin/+ gastrocnemius muscle was reduced 45%, and soleus mean fiber cross-sectional area decreased 24% vs. C57BL/6 mice. Soleus muscle morphology demonstrated pathology of myofibers undergoing degeneration and/or regeneration. These data demonstrate that the ApcMin/+ mouse becomes cachectic by 6 mo of age and that skeletal muscle degeneration and regeneration may be related to the muscle loss.

muscle wasting; colon cancer; inflammatory cytokines

CACHEXIA IS CHARACTERIZED as an overall state of ill health, accompanied by a loss of lean body mass and fat mass, weakness, fatigue, anemia, metabolic abnormalities, inflammation, and impaired immune function, with or without anorexia (4, 5, 38, 45). This condition is associated with other life-threatening diseases, such as cancer and AIDS, especially gastrointestinal tract and lung cancer, and negatively affects life quality and survival (4, 5, 38, 45). Cancer patients can lose up to 30% of their original body weight, and cachexia accounts for ~20–33% of cancer deaths (21). Depressed skeletal muscle function can eventually impair respiratory muscle function, leading to mortality (21, 45). Therefore, elucidating and preventing the events that lead to cachexia may benefit cancer patients by extending life.

The onset and progression of cachexia is multifactorial and has been attributed to increases in reactive oxygen species, metabolic abnormalities, changes in skeletal muscle protein turnover, and systemic inflammation (45). Catalase decreases reactive oxygen species by converting hydrogen peroxide to water and oxygen, and depressed liver catalase activity is a consistent indicator of cachexia (38). Additionally, cachexia is associated with impaired glycolytic enzyme function, decreased ATP synthesis (29), and increased thyroid hormone release (26), which all serve to enhance lipolysis (26, 49) and glucose production (25). In addition to these metabolic abnormalities, skeletal muscle protein synthesis can be decreased (18) and/or protein degradation increased (29). Constituents of the ATP-dependent ubiquitin-proteasome pathway, including 20S proteasome subunits, ubiquitin proteins and ligases, as well as the muscle-specific E3 ligases, atrogin-1/MAFbx and MuRF-I, are upregulated in cachectic muscles (7, 29, 30), which are responsible for both myofiber and extracellular matrix degradation (10, 29). Increased systemic inflammation, which increases circulating IL-6 secretion, can serve to enhance factors related to protein degradation (6, 11, 23, 27, 46), and blockade of this pathway delays or reverses cachexia (46). Additionally, elevated TNF-{alpha} and IFN-{gamma} can selectively decrease myosin heavy chain (MHC) gene expression and increase myofibrillar protein loss in myotubes and mice (1). Inflammation is also associated with degenerating and regenerating myofibers and sustained invasion of inflammatory cells in skeletal muscle that can inhibit the repair process (12, 35, 39, 44). Therefore, tumor-derived reactive oxygen species and inflammatory cytokines may be important for skeletal muscle protein loss and possibly degeneration/regeneration. Impaired skeletal muscle regeneration during cachexia has not been examined.

Tumor transplantation and high TNF-{alpha} or IL-6 cytokine infusion are the most common cachectic rodent models (9, 14, 41). However, these models have several scientific limitations. In tumor transplantation models, the tumor weight may contribute to 30% of the animal's body weight, whereas the human tumor burden is usually <5% of body weight (18). The weight loss in tumor-induced and inflammatory models often has a prominent anorexic component since food intake can be reduced up to 50%. Although anorexia can be associated with cancer, it is often a side effect of chemotherapy (45). Additionally, the duration of these rodent studies typically lasts 2–25 days, whereas in human cachexia, body, muscle, and fat mass decrements occur over months after establishment of the primary disease (1, 6, 41). Inflammatory cytokine dosages administered to rodents to induce wasting do not reflect physiological levels that are attained in human cancer, and these high dosages result in anorexia (19). Although these models have provided valuable initial data elucidating possible mechanisms toward the etiology of cachexia, limitations due to the extreme tumor burden and excessive inflammatory state could mask potential pharmaceutical or nutraceutical agent actions for the prevention or attenuation of muscle wasting.

The ApcMin/+ mouse has been used to elucidate tumor formation and progression related to colon cancer. This mouse may also provide a useful model to study cancer cachexia. The ApcMin/+ mouse has a germline mutation in the adenomatous polyposis coli (Apc) gene, which is responsible for the initiation and development of familial adenomatous polyposis, a type of colon cancer (22). Familial adenomatous polyposis is inherited as an autosomal dominant disorder, and individuals with the disease develop adenomas throughout the colon and rectum. Nonsteroidal anti-inflammatory drugs, such as sulindac and celecoxib, have been shown to be extremely effective for prevention of tumor development and colon cancer progression (37) and have also proven beneficial in the ApcMin/+ mouse (13). This effect is related to cyclooxygenase activity suppression and the inhibition of IL-6 and TNF-{alpha} production. The initial ApcMin/+ tumor burden becomes substantial around 15 wk of age (47), and these mice continue to live for several more months. After 15 wk of age, tumor size continues to increase, which can be influenced by inflammatory cytokines, immune cell infiltration, and increased growth factor expression (24).

A tumor's microenvironment has been described analogous to a wound that fails to heal (16). The tumor stroma continually synthesizes extracellular matrix components and secretes inflammatory chemoattractants that perpetuate the healing process but never fully heal. This serves to maintain the inflammatory microenvironment of the tumor, which can spread systemically. Suppression of protein synthesis and induction of protein degradation have been widely investigated as inflammatory signaling targets for myofiber wasting with cachexia. Systemic inflammation can also initiate muscle necrosis and degeneration, followed by regeneration, characterized by the infiltration of inflammatory cells, centralized nuclei, and the expression of embryonic MHC (eMHC) (35, 39, 44). The incidence of myofiber degeneration and subsequent regeneration in wasting muscle of cachectic animals is not well understood. Therefore, the purpose of the present study was to determine whether ApcMin/+ mice, with a significant intestinal tumor burden, undergo muscle wasting and other systemic characteristics of cachexia, such as muscle degeneration/regeneration. Our primary hypothesis was that muscle wasting would occur in the ApcMin/+ mouse. Our secondary hypothesis was that ApcMin/+ mouse skeletal muscle would exhibit qualities of incomplete regeneration, which would be associated with muscle mass loss.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Animals.   ApcMin/+ male mice on a C57BL/6 background (Jackson Laboratories) were purchased and bred with female C57BL/6 mice in the University of South Carolina's animal resource facility. Offspring were genotyped as heterozygotes by RT-PCR for the Apc gene by taking tail snips at weaning. The primer sequences were sense 5'-TGAGAAAGACAGAAGTTA-3' and antisense 5'-TTCCACTTTGGCATAAGGC-3' (47). The room was maintained on a 16:8-h light-dark cycle with the light period starting at 0500. Mice were provided standard rodent chow (Harlan Teklad Rodent Diet, no. 8604) and water ad libitum. ApcMin/+ female retired breeders (n = 7) and C57BL/6 mice of 6–8 mo of age (n = 7) were housed individually for 3–7 days to measure food and water consumption. Three-month-old ApcMin/+ (n = 7) and C57BL/6 mice (n = 4) were also used. All animal experimentation was approved by the University of South Carolina's Institutional Animal Care and Use Committee.

Tissue collection.   Mice were given a subcutaneous injection of ketamine-xylazine-acepromazine cocktail (1.4 ml/kg body wt). Skeletal muscles, spleens, livers, and tibias were excised. The left soleus was placed in optimal cutting temperature and frozen in isopentane cooled in liquid nitrogen. All other tissues were rinsed in PBS, snap frozen in liquid nitrogen, and stored at –80°C until further analysis. At the time of death, the small and large intestines were removed and qualitatively examined for the presence of tumors. All animals in the study had visible tumors.

Liver catalase activity.   Catalase activity was measured using a procedure modified from Aebi (2). Briefly, frozen liver was homogenized to a final dilution of 1:500 in 100 mM potassium phosphate buffer with 0.05% BSA (pH 7.4). All samples were centrifuged at 2,500 rpm and 4°C for 10 min. To a 1-ml aliquot of supernatant, 200-proof ethanol was added (1:10), vortexed, and incubated on ice for 30 min. Triton X-100 (1% in 100 mM potassium phosphate buffer, no BSA) was added to the ethanol + supernatant reaction (1:10) followed by vortexing and a 15-min incubation on ice. Samples were read in a spectrophotometer for 3 min (25°C, 240 nm). The reaction was started when 150 µl of homogenate mixture was added to 1,050 µl of 10 mM hydrogen peroxide (in 100 mM potassium phosphate buffer, no BSA). Catalase activity was determined by calculating a rate constant (K) = (2.3/t)·[log10(A1/A2)], where t is the time in minutes, A1 is the initial absorbance minus blank absorbance, and A2 is the final absorbance minus blank absorbance. Catalase activity (U/g wet wt) = K·dilution factor, where the dilution factor includes the dilution of the tissue homogenate and the addition of ethanol and Triton X-100. The Bradford assay for protein determination (8) was performed to normalize catalase activity levels to protein concentration.

Circulating IL-6 and TNF-{alpha} measurement.   Plasma IL-6 and TNF-{alpha} were measured using mouse-specific ELISA kits purchased from Biosource as previously described (32). The assays were carried out according to manufacturer's instructions. All samples were run in duplicate, and the interassay coefficients of variation for the IL-6 and TNF-{alpha} assays were 6.7 and 4.6%, respectively.

Total RNA isolation and cDNA synthesis.   Total RNA was isolated using TRIzol reagent (Invitrogen) as previously described (31). Extracted RNA was treated with 10 units of DNase I to degrade any residual contaminating genomic DNA. cDNA was reverse transcribed from 3 µg of total RNA using 1 µl of random hexamers and 50 units of Superscript II reverse transcriptase (Invitrogen) in a final volume of 20 µl at 25°C for 10 min, followed by 42°C for 50 min and 70°C for 15 min.

Semiquantitative PCR.   PCR amplication was performed as previously described (31). Primers for IL-6 (40), TNF-{alpha} (17), and 18S (48) were synthesized by Integrated DNA Technologies (Caralville, IA) containing the following base sequences: IL-6: sense, 5'-GCCTATTGAAAATTTCCTCTG-3', antisense, 5'-GTTTGCCGAGTAGATCTC-3'; TNF-{alpha}: sense, 5'-CCCAGACCCTCACACTCAGAT-3', antisense, 5'-TTGTCCCTTGAAGAGAACCTG-3'; 18S: sense, 5'-AAACGGCTACCACATCCAAG-3', antisense, 5'-CCCTCTTAATCATGGCCTCA-3'. The number of cycles and the PCR conditions for each target mRNA were optimized so that the amplified signal was still on the linear portion of the amplification curve. PCR started with an initial denaturation at 95°C for 10 min. Amplification was carried out by repeating the following cycles for TNF-{alpha} (36 cycles), IL-6 (39 cycles), and 18S (16 cycles): denaturation at 94°C for 45 s, annealing at 60°C, and extension at 72°C for 60 s, with a final extension cycle at 72°C for 10 min. Amplified products were subjected to electrophoresis through 2.0% agarose gels, stained with ethidium bromide, visualized by ultraviolet transillumination, and photographed, and then quantified by densitometry scanning (Scion Image, Frederick, MD). Target fragment levels were normalized against 18S, and the data were presented as each cytokine mRNA integrated optical density-to-18S integrated optical density ratio.

Total RNA and protein measurement.   Total RNA was quantified in the spleen and gastrocnemius by using the method of Fleck and Munro (20). Briefly, a frozen tissue (~20 mg) was cut and weighed. The tissue was homogenized in 0.2 N HClO4 and centrifuged (4°C, 12,000 rpm, 10 min). After two washes in 0.2 N HClO4, the pellet was dried and suspended in 0.3 N KOH and incubated at 37°C overnight. An aliquot was removed and used in the Bradford assay for protein determination of the gastrocnemius (8). To the remaining sample, 0.75 ml of 1.2 N HClO4 was added. After centrifugation (4°C, 12,000 rpm, 10 min), the supernatant was transferred to a new tube, the pellet was washed two more times with 0.2 N HClO4, and the subsequent supernatants were pooled from all washes. An aliquot was diluted 1:10 and read on a spectrophotometer at 260 nm to determine total RNA concentration.

Soleus morphological analysis.   Sectioning of muscle and staining are the same as described previously (31). Briefly, transverse sections (10 µm) were cut from the mid-belly of the soleus muscle on a cryostat at –20°C. Hematoxylin and eosin (H & E) staining was performed on sections from all animals to measure fiber cross-sectional area and regenerating fibers. Digital photographs were taken from each H & E section at a x40 magnification with a Kodak spot camera, and ~100 fibers/animal were traced with imaging software (Scion Image) in a blinded fashion. Nuclei were also counted on these images and categorized as myofiber or extracellular matrix nuclei. The criteria for central nuclei were defined as normal fibers that contained a well-defined nucleus, equidistant from the surrounding sarcolemma. The area of the extracellular matrix was determined by overlaying an 18 x 14 digital pixel grid and counting which pixels were associated with a myofiber or with the extracellular matrix. Pixels were counted if they covered at least 75% of the designated area. Pixels that were indistinguishable were omitted from the total count.

Immunohistochemistry.   Frozen sections of soleus were cut on a cryostat at 10-µm sections. Sections were air-dried and fixed in acetone for 10 min. Samples were quenched in 0.3% H2O2 for 2 h and blocked in 4% horse serum for 3 h. eMHC (University of Iowa Hybridoma Bank) was added to each sample (1:10) in PBS. Slides were washed in PBS and incubated with anti-mouse IgG secondary (1:200) in PBS and 1.5% horse serum. Color detection was visualized with an avidin-biotin complex detection kit and diaminobenzidine (Vector). Digital photographs were taken from each sample at a x40 magnification with a Kodak spot camera. Fibers that stained positive for eHMC were counted by an individual blinded to the treatment and expressed as the number of eMHC-positive fibers per mm2.

Fiber-type identification.   Soleus sections were cut on a cryostat at 10-µm sections and were reacted for ATPase preincubated at pH 9.8 (8). Soleus muscle fibers were classified as type I (no staining), type IIa (dark staining), or type IIb (light staining). Digital images were taken at a x20 magnification, and ~300 fibers per animal were classified as type I, type IIa, or type IIb and expressed as a percentage of the total fiber distribution.

Western blot analysis.   Western blotting was performed as described previously (31). Briefly, frozen gastrocnemius muscle was homogenized in Mueller buffer, and protein concentration was determined by the Bradford method (8). Crude muscle homogenate (5–60 µg) was fractionated on 8–12% polyacrylamide gels. Gels were transferred to polyvinylidene difluoride membranes overnight and blocked in 5% TBS-Tween milk. Equal protein loading of the gels was qualitatively assessed using Ponceau staining. Primary antibodies for the 20s proteasome {alpha}/{beta} subunits (BIOMOL International), phosphorylated p70s6kinase at Thr 389 (Cell Signaling Technology), and {beta}-catenin (Cell Signaling Technology) were incubated at a 1:1,000 dilution for 1 h in 1% TBS-Tween milk. Secondary anti-rabbit IgG conjugated secondary antibodies (Amersham Biosciences) were incubated with the membranes at a 1:5,000 dilution for 1 h in 1% TBS-Tween milk. Enhanced chemiluminescence (Amersham Biosciences) was used to visualize the antibody-antigen interactions and developed by autoradiography (Kodak, Biomax). Digitally scanned blots were analyzed by measuring the integrated optical density of each band using digital imaging software (Scion Image).

Statistical analysis.   Results are reported as means ± SE. Data were analyzed with a two-way ANOVA to determine age and strain effects for body weight, muscle weights, spleen weight, tibia length, and liver catalase activity. Student-Newman-Keuls post hoc comparisons were made when an interaction existed. All other variables were analyzed with Student's t-tests. P ≤ 0.05 was accepted as the level of statistical significance.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Inflammatory state.   Cancer is often accompanied by a decrease in antioxidant defenses and an increase in systemic inflammation due to the tumor burden. Although the tumor burden was not quantified, all intestines were qualitatively examined and multiple adenomas were detected in all ApcMin/+ mice. Additionally, mice by this age typically have >100 intestinal polyps (47). To confirm the diseased state of 6-mo-old ApcMin/+ mice, liver catalase activity was measured. Liver catalase activity was 54% less in 6-mo-old ApcMin/+ mice compared with C57BL/6 mice (P < 0.001) (Fig. 1A). Splenomegaly, which increases during inflammation, and inflammatory cytokine expression were examined in 6-mo-old C57BL/6 and ApcMin/+ mice. ApcMin/+ mouse spleen weight was increased 329% compared with C57BL/6 mice (P < 0.001) (Fig. 1B). ApcMin/+ mouse splenic IL-6 (P = 0.009) and TNF-{alpha} (P = 0.034) mRNA concentrations were both elevated 2.1-fold over C57BL/6 mice. Plasma IL-6 was not detected in C57BL/6 mice but was elevated in ApcMin/+ mice (P = 0.028) (Fig. 1C). However, plasma TNF-{alpha} was not different between ApcMin/+ (18.2 ± 13.0 pg/ml) and C57BL/6 mice (15.4 ± 8.0 pg/ml).



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  		Fig. 1. Plasma IL-6 and liver catalase activity in 6-mo-old female C57BL/6 and ApcMin/+ mice. A: plasma IL-6. B: liver catalase activity. Data were analyzed with independent t-tests. Significance was set at P < 0.05. *Significant difference from C57BL/6 (P < 0.05).

 
Animal characteristics.   At 6 mo of age, ApcMin/+ mice weighed 21% less than C57BL/6 mice (P < 0.001) (Table 1). Despite the difference in body mass, this was not due to anorexia since C57BL/6 and ApcMin/+ mice consumed similar amounts of food (4.4 ± 0.2 vs. 4.9 ± 0.7 g/day; P = 0.562). Part of the differences in body weight can be attributed to skeletal muscle loss in the ApcMin/+ mice. The gastrocnemius muscle wet weight decreased 45% in ApcMin/+ mice compared with C57BL/6 mice (P < 0.001). Soleus muscle wet weight was 25% less in ApcMin/+ mice compared with C57BL/6 mice (P < 0.001). It should also be noted that the ApcMin/+ mouse plantaris (14 ± 1 vs. 10 ± 1 mg; P = 0.006) and tibialis anterior (42 ± 3 vs. 27 ± 5 mg; P = 0.034) muscles also demonstrated significant atrophy.


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  		Table 1. Body weight, hindlimb skeletal muscle wet weights, and muscle-to-tibia length ratios from 3- and 6-mo female C57BL/6 and ApcMin/+ mice

 
ApcMin/+ mouse development.   To determine whether ApcMin/+ mice had abnormal skeletal muscle development and growth, 3-mo-old animals were also examined. At 3 mo of age, there were no differences in body weight between C57BL/6 and ApcMin/+ mice (P = 0.614) (Table 1). In addition, C57BL/6 and ApcMin/+ soleus (P = 0.897) and gastrocnemius muscles (P = 0.728) were similar. To determine whether the muscle mass decrease in 6-mo-old ApcMin/+ mice was due to growth inhibition rather than atrophy, tibia length was used as an indicator of animal size and growth. Although there was a significant main effect of age on tibia length (P < 0.001), there were no differences between C57BL/6 and ApcMin/+ mice at 3 or 6 mo of age (P = 0.761). The soleus and gastrocnemius were also decreased in 6-mo-old ApcMin/+ mice when normalized to tibia length. In addition to bone growth, body weight increased in both C57BL/6 (P < 0.001) and ApcMin/+ mice (P = 0.047) from 3 to 6 mo of age. Despite an increase in body weight from 3 to 6 mo of age, the ApcMin/+ gastrocnemius muscle decreased in size (P = 0.004), whereas the C57BL/6 gastrocnemius muscle increased in size (P < 0.001). The ApcMin/+ soleus did not change mass from 3 to 6 mo of age (P = 0.414), but the C57BL/6 soleus did increase mass during this time (P = 0.008). Because the ApcMin/+ mouse is a heterozygote for the Apc gene, it is possible that this mutation might affect skeletal muscle Wnt->APC->{beta}-catenin signaling pathway, which is implicated in skeletal muscle development. However, 6-mo-old ApcMin/+ and C57BL/6 mice expressed similar amounts of gastrocnemius {beta}-catenin protein (Fig. 2).



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  		Fig. 2. Gastrocnemius {beta}-catenin protein levels in 6-mo-old C57BL/6 and ApcMin/+ mice.

 
Gastrocnemius protein loss.   To determine the protein synthetic capacity of the ApcMin/+ skeletal muscle, the gastrocnemius muscle was used to analyze total muscle RNA and protein. Protein and RNA concentrations were not significantly different between 6-mo-old C57BL/6 and ApcMin/+ mice. However, due to the differences in wet muscle weights, total gastrocnemius muscle protein (P < 0.001) and RNA (P < 0.001) were 44 and 37% lower, respectively, in 6-mo-old ApcMin/+ mice compared with C57BL/6 mice (Table 2). Western blotting was performed to determine the relative amounts of phosphorylated p70s6kinase, a marker of skeletal muscle growth and protein synthesis, and the 20S proteasome subunit, a marker of protein degradation. There were no significant changes in the expression of phosphorylated p70s6kinase or the 20S proteasome subunits in the gastrocnemius muscle of 6-mo-old C57BL/6 and ApcMin/+ mice.


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  		Table 2. Gastrocnemius mass atrophy does not alter expression of the 20S proteasome or p70s6 kinase, phosphorylation in 6-mo-old ApcMin/+ mice

 
Soleus morphology.   To determine whether the skeletal muscle loss was due to myofiber atrophy, soleus fiber cross-sectional area and extracellular matrix area were analyzed on soleus H & E stained sections from 6-mo-old C57BL/6 and ApcMin/+ mice (see Fig. 5, A and B). Soleus muscle mean fiber cross-sectional area decreased 24% in ApcMin/+ compared with C57BL/6 mice (P = 0.001) (Fig. 3A). There were no differences in the percentage of the extracellular matrix between C57BL/6 and ApcMin/+ mice (P = 0.770) (Fig. 3B). Fiber-type switching also occurs during different models of skeletal muscle atrophy. ATPase staining was performed to determine the fiber-type distribution of the soleus muscle (see Fig. 5, C and D). There was a 15% reduction in the percentage of type I fibers (P = 0.013) that accompanied a 15% increase in type IIa fibers (P = 0.028) and the expression of type IIb fibers in 6-mo-old ApcMin/+ mice (Fig. 3C). Three-month-old soleus fiber distribution was also examined, and there were no differences in the percentage of type I, IIa, or IIb fibers between the two strains.



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  		Fig. 5. Representative images demonstrating a soleus muscle fiber type, fiber area, central nuclei, extracellular nuclei, and eMHC-positive staining in 6-mo-old female C57BL/6 and ApcMin/+ mice. A: C57BL/6 ATPase staining (10x). B: ApcMin/+ ATPase staining (10x). C: C57BL/6 hematoxylin and eosin (40x). D: ApcMin/+ hematoxylin and eosin (40x). E: C57BL/6 eMHC (40x). F: ApcMin/+ eMHC (40x).

 


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  		Fig. 3. Soleus muscle fiber cross-sectional area (CSA; A), extracellular matrix (ECM) area (B), and fiber-type distribution (C) in 6-mo-old female C57BL/6 and ApcMin/+ mice. Hematoxylin and eosin sections were stained, and x40 images were analyzed using imaging software to determine fiber CSA and ECM area. ATPase staining (10x images) was used to determine type I, type IIa, and type IIb fibers for fiber-type distribution. *Significant difference from C57BL/6 (P < 0.05).

 
Soleus degeneration/regeneration.   Damaging stimuli to skeletal muscle initiate a period of myofiber degeneration classified by centrally located nuclei and infiltrating immune cells. To determine whether the ApcMin/+ mouse exhibits signs of degeneration and/or regeneration, C57BL/6 and ApcMin/+ mouse soleus nuclei were counted on 40x H & E stained sections. Nuclei were classified as associated with the myofiber, extracellular matrix, or centralized nuclei. There was no difference in the number of myofiber nuclei between C57BL/6 (548 ± 62 nuclei/mm2) and ApcMin/+ mice (497 ± 65 nuclei/mm2; P = 0.583). However, there was a significant 17% increase in total nuclei in ApcMin/+ (1,811 ± 101 nuclei/mm2) compared with C57BL/6 mice (1,545 ± 55 nuclei/mm2; P = 0.042), which was due to a 33% increase in extracellular matrix nuclei (P = 0.034; Fig. 4A). The number of centralized nuclei (Fig. 4B) increased threefold in ApcMin/+ compared with C57BL/6 mice (P = 0.039). These data were confirmed by immunohistochemical detection of eMHC (Fig. 5, E and F). The number of eMHC-positive fibers increased 12-fold in ApcMin/+ compared with C57BL/6 mice (Fig. 4C).



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  		Fig. 4. Regenerative characteristics of 6-mo-old female C57BL/6 and ApcMin/+ soleus muscle. Hematoxylin and eosin sections were stained and 40x pictures were analyzed using imaging software to determine nuclear localization. Immunohistochemistry was used to determine embryonic myosin heavy chain (eMHC) positive fibers. A: soleus ECM nuclei. B: soleus central nuclei. C: soleus eMHC-positive cells. Data were analyzed with independent t-tests. Significance was set at P < 0.05. *Significant difference from C57BL/6 (P < 0.05).

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Systemic inflammation during cancer can initiate and propagate skeletal muscle wasting through altered protein turnover (1, 29), but it is not known how induction of inflammatory-mediated myofiber degeneration contributes to muscle loss during cachexia. The present study's major finding is that female ApcMin/+ mice undergo muscle mass loss by 6 mo of age, which is associated with extensive evidence of degeneration and regeneration in the slow-type soleus muscle. The inability to complete the regenerative process in the presence of chronic inflammation may be responsible for a portion of the ApcMin/+ mouse skeletal muscle wasting. Previous studies examining mechanisms inducing this wasting environment have utilized animal models that manipulate either cancer severity by tumor implantation or inflammatory stress by cytokine infusion (9, 14, 41). To our knowledge, the present study is the first demonstrating cachexia in the ApcMin/+ mouse, a genetic model of cancer with a significant intestinal polyp burden.

The initial purpose of this study was to determine whether ApcMin/+ mice were exhibiting classical systemic characteristics that are associated with cachexia. Body weight loss is a hallmark of cachexia and is a significant predictor of mortality in cancer patients and comprises both lean body mass and fat mass (45). Tumor-bearing animals can decrease body weight 20% within 3 wk (1, 14), whereas in humans, weight loss ranges from 5 to 30% over longer time periods (45). ApcMin/+ mice weighed ~20% less than age-matched wild-type mice, which is similar to both human and other animal models of cachexia. ApcMin/+ weight loss did not appear until after 3 mo of age, which is after animal tumor burden has been shown to be substantial (47). Several lines of evidence suggest body mass loss was not due to altered growth in the ApcMin/+ mouse. Wild-type and ApcMin/+ mice had similar body weights and tibia lengths at 3 mo of age. Bone growth continued from 3 to 6 mo of age regardless of strain, and tibia lengths remained similar between the two strains over this time period. Unlike bone growth, ApcMin/+ gastrocnemius muscle mass actually decreased between 3 and 6 mo of age, whereas wild-type muscle mass increased over this time. Malnutrition or anorexia are possible explanations for the body mass loss. Lifetime nutritional intervention studies have demonstrated that nutrient absorption is not limited by tumor burden in ApcMin/+ mice (34). Anorexia is associated with both human and animal models of cachexia but typically cannot account for the total body mass loss (45). Inadequate caloric consumption or starvation also initially results in fat mass loss, whereas lean body mass is preserved (45). The ApcMin/+ mouse body mass loss was independent of anorexia since ApcMin/+ and wild-type mice ate similarly. In fact, food intake per body weight would actually be increased in the ApcMin/+ mouse. ApcMin/+ mice also showed a reduction in skeletal muscle mass, which is uncharacteristic of anorexia.

Skeletal muscle mass loss impairs muscle function and is linked to frailty, morbidity, and mortality (45). Skeletal muscle atrophy can be induced by a variety of stimuli including aging, disuse, diabetes, sepsis, and cancer. However, the rate and type of skeletal muscle loss can differ depending on the atrophic stimulus (29). Muscle atrophy related to cancer cachexia ranges from 14 to 45% loss, and fast-twitch fibers have been reported to be more susceptible to atrophy compared with slow-oxidative fibers (1, 15). MHC type IIa and IIb are preferentially targeted for degradation by circulating TNF-{alpha} and IL-6 during cancer cachexia, whereas components of myosin light chains and type I MHC are unaffected (1). The gastrocnemius muscle is composed primarily of type II fibers and the ApcMin/+ mouse gastrocnemius muscle mass and total protein content were reduced 45 and 44%, respectively, compared with wild-type mice. The soleus, a highly oxidative postural muscle, atrophied only 25%, with a corresponding 24% reduction in mean fiber cross-sectional area. Since type II MHC can be degraded during wasting conditions, whereas type I MHCs remain unaffected, the larger amount of muscle loss in the gastrocnemius relative to the soleus might be related to the fiber-type composition of each skeletal muscle. The preferential loss of type II fibers was not seen in the ApcMin+ soleus muscle. The 6-mo-old ApcMin/+ soleus muscle exhibited an increase in type IIa and IIb fibers, which has previously been reported in cachectic mice (15). Diffee et al. (15) suggested that the shift toward a faster phenotype during cachexia is tumor dependent, most likely related to proinflammatory cytokines and not related to anorexia or activity level, which can also alter the myosin phenotype. The stimulus for the phenotype shift in the present study is not clear, since both systemic inflammation and reduced activity were likely present. Disuse atrophy typically induces a shift toward a faster soleus phenotype (3). The ApcMin/+ mouse may decrease ambulatory activity with age. Studies in our laboratory have demonstrated that ApcMin/+ mice decrease voluntary wheel running activity and develop anemia after 14 wk of age and continue to progressively decline until 6 mo of age (data not shown). Therefore, the shift in soleus fiber-type profile may be a reflection of the decreased ambulatory activity of the animal rather than factors related to chronic inflammation's effect on skeletal muscle. Further work is needed to delineate whether the pattern of soleus myosin isoform expression is caused by inactivity or activation of signaling pathways induced during skeletal muscle wasting.

Inflammatory cytokines are potential mediators of skeletal muscle wasting, and excess production of TNF-{alpha}, IL-6, and IL-1 can alter protein turnover (5, 38, 45). IL-6 abundance by genetic manipulation, tumor implantation, or local infusion produces a significant reduction in skeletal muscle mass and protein content (6, 11, 23, 27, 46). Inflammatory cytokines can also inhibit extracellular matrix protein synthesis (10). Inflammatory cytokines are produced not only from tumors during cancer but can also mediate cytokine synthesis in other organs and tissues, such as the spleen (6). Furthermore, muscle loss during cachexia can be reversed by anti-cytokine therapies using IL-6-neutralizing antibodies, drugs to block the action at the receptor, or interfering with related inflammatory pathways (14, 43). Increased circulating IL-6 levels, splenomegaly, and increased splenic TNF-{alpha} and IL-6 gene expression all demonstrate that the ApcMin/+ mouse was chronically inflamed. Although TNF-{alpha} has been implicated in wasting syndromes, circulating levels were not induced and suggest that IL-6 may be a more critical mediator of the chronic inflammation and muscle wasting in the ApcMin/+ mouse. However, tissue-specific TNF-{alpha} production and signaling in muscle could be independent of systemic levels and require further study.

Increased protein degradation can be accomplished via the ubiquitin-proteasome pathway, the calcium-dependent pathway, or the lysosomal pathway. Components of the ATP-dependent ubiquitin-proteasome pathway are typically upregulated during cachexia (7, 29, 45). Protein degradation, as measured by the abundance of the 20S proteasome, was not elevated in the ApcMin/+ mouse gastrocnemius muscle. A lack of increase in the 20S proteasome subunits does not rule out ubiquitin-proteasome regulation of protein degradation. The ubiquitin-proteasome pathway is complex, and overall activity of the pathway has been shown to change independent of individual components (36). Furthermore, it is also possible that other components of the ubiquitin-proteasome pathway were elevated, such as the level of ubiquitinated proteins, E2-conjugating enzymes, or E3 ligases, particularly the muscle-specific E3 ligases, MuRF-I or atrogin-I/MAFbx. Additionally, the slower rate of wasting in the ApcMin/+ mouse compared with other models may impact ubiquitin-proteasome regulation. Wasting in the present study occurred over a period of 3 mo. During muscle disuse, such as immobilization, denervation, and hindlimb suspension, the expression of MuRF-1, MAFbx, and proteasome subunit mRNAs increases during initial muscle loss (7, 29, 36). However, expression of these atrophy genes begins to return to baseline levels during longer periods of disuse, despite more atrophy (7, 36). Therefore, it is possible that elevated expression of the ApcMin/+ gastrocnemius 20S proteasome subunits occurred during the beginning of the wasting process and had reached equilibrium by 6 mo of age. Additionally, the reported stimulation of this protein degradation pathway with wasting has occurred in conditions with extremely high levels of IL-6 and were accompanied by an extremely rapid muscle mass loss (1, 29, 45). Mice in the present study had chronic low levels of elevated IL-6, up to 30 times lower than those previously reported for chronically administered IL-6 (27). Corresponding with the chronic low level of inflammation was a gradual loss of muscle mass occurring between 3 and 6 mo of age. The slower rate of muscle loss may have more subtle changes in muscle protein turnover. Further work is needed to determine whether the calcium-dependent or lysosomal-mediated proteolysis may be critical for muscle mass loss in the ApcMin/+ mouse.

Inhibition of skeletal muscle protein synthesis has also been implicated during cancer cachexia and can be mediated by inflammatory pathways (18, 23, 29, 45). IGF-I is a critical mediator of skeletal muscle growth and hypertrophy. During skeletal muscle hypertrophy, IGF-I binds to its transmembrane receptor and activates a series of kinases, including Akt. Downstream targets of Akt include p70s6kinase, glycogen synthase kinase, and mammalian target of rapamycin, all of which are proteins involved in protein translation and synthesis. During skeletal muscle wasting, activation of Akt signaling can become impaired and skeletal muscle mass restoration can be achieved by IGF-I or Akt overexpression (23, 28). Phosphorylation of p70s6kinase is decreased during skeletal muscle wasting and is associated with the upregulation of IL-6 (23, 30, 42). In the present study, p70s6kinase phosphorylation was not depressed in the ApcMin/+ mouse gastrocnemius muscle. It is possible that decreases in p70s6kinase phosphorylation and other protein synthetic pathways are present early in the ApcMin/+ cachectic process but reach equilibrium by 6 mo of age. Additionally, since total muscle RNA concentrations were similar between the two strains, ApcMin/+ gastrocnemius muscle translational capacity did not appear to be impaired. Nutritional status can also affect this signaling pathway, and food intake did not differ between the strains, as mentioned previously. Although these data suggest that the mechanisms leading to ApcMin/+ skeletal muscle wasting are not related to protein synthesis inhibition, earlier time points of ApcMin/+ skeletal muscle wasting need further investigation.

The intricate balance of protein synthesis and protein degradation is important during wound healing and skeletal muscle regeneration. Tumors have been described as wounds that fail to heal, and this environment may promote similar effects on skeletal muscle during cachexia (16, 33). Skeletal muscle consists of postmitotic, multinucleated fibers that are capable of self-regeneration during damaging stimuli (12). During the initial phases of skeletal muscle regeneration, muscle fibers exhibit sarcolemma damage driven by calcium-dependent proteolysis, infiltration of mononucleated cells, and small fibers with centrally located nuclei that express eMHC (12). Interestingly, tumor invasion of skeletal muscle results in similar alterations, such as ED-1 macrophage infiltration, centronucleation, and sparse myofibers (33). The ApcMin/+ mouse soleus muscle displayed an increase in centralized nuclei, eMHC-positive cells, and the infiltration of extracellular matrix nuclei, similar to the initial phase of skeletal muscle regeneration. Mononucleated cells include macrophages and neutrophils, which are responsible for the phagocytic removal of damaged tissue but can also mediate cell damage by releasing free radicals (44). Because the ApcMin/+ mouse soleus muscle exhibits an increase in the number of extracellular matrix nuclei, these cells could be inflammatory cells initiating cellular damage. Alterations in the immune cell response can delay skeletal muscle regeneration (44). Because calcium-dependent proteolysis is responsible for the necrosis during the initial phases of skeletal muscle repair, this also suggests that ApcMin/+ skeletal muscle loss may be functioning through a calpain-dependent mechanism rather than the ATP-dependent ubiquitin-proteasome pathway. This may keep the muscle in a constant state of attempting to heal, which is associated with degeneration and necrosis, ultimately leading to muscle loss.

In summary, the tumor burden of the ApcMin/+ mouse creates an environment that promotes skeletal muscle atrophy and degeneration and/or regeneration. ApcMin/+ mouse skeletal muscle atrophy affects both the myofibers and connective tissue. The capacity for protein synthesis is not altered, but the soleus muscle appears to be in a constant state of regeneration. Centralized nuclei, eMHC-expressing fibers, and the infiltration of extracellular nuclei demonstrate that ApcMin/+ mouse soleus muscle is in a state of repair. Potential therapies that target the regeneration process may provide relief to those with muscle-wasting diseases.


   GRANTS
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 ABSTRACT
 METHODS
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 DISCUSSION
 GRANTS
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The research described in this report was supported by National Center for Research Resources Grant P20 RR-017698. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the National Institutes of Health.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
The authors thank Tia Davis and Julie Clements for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. A. Carson, Univ. of South Carolina, Dept. of Exercise Science, 1300 Wheat St., Columbia, SC 29208 (e-mail: carsonj{at}sc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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  1. Acharyya S, Ladner KJ, Nelsen LL, Damrauer J, Reiser PJ, Swoap S, and Guttridge DC. Cancer cachexia is regulated by selective targeting of skeletal muscle gene products. J Clin Invest 114: 370–378, 2004.[CrossRef][Web of Science][Medline]
  2. Aebi H. Catalase in vitro. Methods Enzymol 105: 121–126, 1984.[Web of Science][Medline]
  3. Agbulut O, Noirez P, Beaumont F, and Butler-Browne G. Myosin heavy chain isoforms in postnatal muscle development of mice. Biol Cell 95: 399–406, 2003.[CrossRef][Web of Science][Medline]
  4. Ardies CM. Exercise, cachexia, and cancer therapy: a molecular rationale. Nutr Cancer 42: 143–157, 2002.[CrossRef][Web of Science][Medline]
  5. Argiles JM, Busquets S, and Lopez-Soriano FJ. Cytokines in the pathogenesis of cancer cachexia. Curr Opin Clin Nutr Metab Care 6: 401–406, 2003.[CrossRef][Web of Science][Medline]
  6. Barton BE and Murphy TF. Cancer cachexia is mediated in part by the induction of IL-6-like cytokines from the spleen. Cytokine 16: 251–257, 2001.[CrossRef][Web of Science][Medline]
  7. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela DM, DeChiara TM, Stitt TN, Yancopoulos GD, and Glass DJ. Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294: 1704–1708, 2001.[Abstract/Free Full Text]
  8. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248–254, 1976.[CrossRef][Web of Science][Medline]
  9. Buck M and Chojkier M. Muscle wasting and dedifferentiation induced by oxidative stress in a murine model of cachexia is prevented by inhibitors of nitric oxide synthesis and antioxidants. EMBO J 15: 1753–1765, 1996.[Web of Science][Medline]
  10. Buck M, Houglum K, and Chojkier M. Tumor necrosis factor-alpha inhibits collagen alpha1(I) gene expression and wound healing in a murine model of cachexia. Am J Pathol 149: 195–204, 1996.[Abstract]
  11. Cai D, Frantz JD, Tawa NE Jr, Melendez PA, Oh BC, Lidov HG, Hasselgren PO, Frontera WR, Lee J, Glass DJ, and Shoelson SE. IKK{beta}/NF-{kappa}B activation causes severe muscle wasting in mice. Cell 119: 285–298, 2004.[CrossRef][Web of Science][Medline]
  12. Charge SB and Rudnicki MA. Cellular and molecular regulation of muscle regeneration. Physiol Rev 84: 209–238, 2004.[Abstract/Free Full Text]
  13. Corpet DE and Pierre F. Point: from animal models to prevention of colon cancer. Systematic review of chemoprevention in min mice and choice of the model system. Cancer Epidemiol Biomarkers Prev 12: 391–400, 2003.[Abstract/Free Full Text]
  14. Davis TW, Zweifel BS, O'Neal JM, Heuvelman DM, Abegg AL, Hendrich TO, and Masferrer JL. Inhibition of cyclooxygenase-2 by celecoxib reverses tumor-induced wasting. J Pharmacol Exp Ther 308: 929–934, 2004.[Abstract/Free Full Text]
  15. Diffee GM, Kalfas K, Al-Majid S, and McCarthy DO. Altered expression of skeletal muscle myosin isoforms in cancer cachexia. Am J Physiol Cell Physiol 283: C1376–C1382, 2002.[Abstract/Free Full Text]
  16. Dvorak HF. Tumors: wounds that do not heal. Similarities between tumor stroma generation and wound healing. N Engl J Med 315: 1650–1659, 1986.[Web of Science][Medline]
  17. Elliott JL. Cytokine upregulation in a murine model of familial amyotrophic lateral sclerosis. Brain Res Mol Brain Res 95: 172–178, 2001.[Medline]
  18. Emery PW, Lovell L, and Rennie MJ. Protein synthesis measured in vivo in muscle and liver of cachectic tumor-bearing mice. Cancer Res 44: 2779–2784, 1984.[Abstract/Free Full Text]
  19. Finck BN and Johnson RW. Anorexia, weight loss and increased plasma interleukin-6 caused by chronic intracerebroventricular infusion of interleukin-1{beta} in the rat. Brain Res 761: 333–337, 1997.[CrossRef][Web of Science][Medline]
  20. Fleck A and Munro HN. The precision of ultraviolet absorption measurements in the Schmidt-Thannhauser procedure for nucleic acid estimation. Biochim Biophys Acta 55: 571–583, 1962.[Medline]
  21. Giordano A, Calvani M, Petillo O, Carteni M, Melone MR, and Peluso G. Skeletal muscle metabolism in physiology and in cancer disease. J Cell Biochem 90: 170–186, 2003.[CrossRef][Web of Science][Medline]
  22. Groden J, Thliveris A, Samowitz W, Carlson M, Gelbert L, Albertsen H, Joslyn G, Stevens J, Spirio L, Robertson M, et al. Identification and characterization of the familial adenomatous polyposis coli gene. Cell 66: 589–600, 1991.[CrossRef][Web of Science][Medline]
  23. Haddad F, Zaldivar F, Cooper DM, and Adams GR. IL-6-induced skeletal muscle atrophy. J Appl Physiol 98: 911–917, 2005.[Abstract/Free Full Text]
  24. Hanahan D and Weinberg RA. The hallmarks of cancer. Cell 100: 57–70, 2000.[CrossRef][Web of Science][Medline]
  25. Holroyde CP, Skutches CL, Boden G, and Reichard GA. Glucose metabolism in cachectic patients with colorectal cancer. Cancer Res 44: 5910–5913, 1984.[Abstract/Free Full Text]
  26. Hyltander A, Drott C, Korner U, Sandstrom R, and Lundholm K. Elevated energy expenditure in cancer patients with solid tumours. Eur J Cancer 27: 9–15, 1991.[Web of Science][Medline]
  27. Janssen SP, Gayan-Ramirez G, Van den Bergh A, Herijgers P, Maes K, Verbeken E, and Decramer M. Interleukin-6 causes myocardial failure and skeletal muscle atrophy in rats. Circulation 111: 996–1005, 2005.[Abstract/Free Full Text]
  28. Lai KM, Gonzalez M, Poueymirou WT, Kline WO, Na E, Zlotchenko E, Stitt TN, Economides AN, Yancopoulos GD, and Glass DJ. Conditional activation of akt in adult skeletal muscle induces rapid hypertrophy. Mol Cell Biol 24: 9295–9304, 2004.[Abstract/Free Full Text]
  29. Lecker SH, Jagoe RT, Gilbert A, Gomes M, Baracos V, Bailey J, Price SR, Mitch WE, and Goldberg AL. Multiple types of skeletal muscle atrophy involve a common program of changes in gene expression. FASEB J 18: 39–51, 2004.[Abstract/Free Full Text]
  30. McCarthy DO, Whitney P, Hitt A, and Al-Majid S. Indomethacin and ibuprofen preserve gastrocnemius muscle mass in mice bearing the colon-26 adenocarcinoma. Res Nurs Health 27: 174–184, 2004.[CrossRef][Web of Science][Medline]
  31. McClung JM, Lee WJ, Thompson RW, Lowe LL, and Carson JA. RhoA induction by functional overload and nandrolone decanoate administration in rat skeletal muscle. Pflügers Arch 447: 345–355, 2003.[CrossRef][Web of Science][Medline]
  32. Mehl KA, Davis JM, Clements JM, Berger FG, Pena MM, and Carson JA. Decreased intestinal polyp multiplicity is related to exercise mode and gender in ApcMin/+ mice. J Appl Physiol 98: 2219–2225, 2005.[Abstract/Free Full Text]
  33. Ohba N, Maeda M, Sakamoto H, Kiyama H, Ishii M, Muraoka M, and Kaneda K. Skeletal muscle regeneration associated with the stroma reaction during tumor invasion in the rat tongue. J Submicrosc Cytol Pathol 34: 367–376, 2002.[Medline]
  34. Perkins S, Clarke AR, Steward W, and Gescher A. Age-related difference in susceptibility of Apc(Min/+) mice towards the chemopreventive efficacy of dietary aspirin and curcumin. Br J Cancer 88: 1480–1483, 2003.[CrossRef][Web of Science][Medline]
  35. Pizza FX, Peterson JM, Baas JH, and Koh TJ. Neutrophils contribute to muscle injury and impair its resolution after lengthening contractions in mice. J Physiol 562: 899–913, 2005.[Abstract/Free Full Text]
  36. Reid MB. Response of the ubiquitin-proteasome pathway to changes in muscle activity. Am J Physiol Regul Integr Comp Physiol 288: R1423–R1431, 2005.[Abstract/Free Full Text]
  37. Rigas B and Williams JL. NO-releasing NSAIDs and colon cancer chemoprevention: a promising novel approach (Review). Int J Oncol 20: 885–890, 2002.[Web of Science][Medline]
  38. Rubin H. Cancer cachexia: its correlations and causes. Proc Natl Acad Sci USA 100: 5384–5389, 2003.[Abstract/Free Full Text]
  39. Salvadori C, Peters IR, Day MJ, Engvall E, and Shelton GD. Muscle regeneration, inflammation, and connective tissue expansion in canine inflammatory myopathy. Muscle Nerve 31: 192–198, 2005.[CrossRef][Web of Science][Medline]
  40. Sass G, Heinlein S, Agli A, Bang R, Schumann J, and Tiegs G. Cytokine expression in three mouse models of experimental hepatitis. Cytokine 19: 115–120, 2002.[CrossRef][Web of Science][Medline]
  41. Smith HJ, Greenberg NA, and Tisdale MJ. Effect of eicosapentaenoic acid, protein and amino acids on protein synthesis and degradation in skeletal muscle of cachectic mice. Br J Cancer 91: 408–412, 2004.[CrossRef][Web of Science][Medline]
  42. Song YH, Li Y, Du J, Mitch WE, Rosenthal N, and Delafontaine P. Muscle-specific expression of IGF-1 blocks angiotensin II-induced skeletal muscle wasting. J Clin Invest 115: 451–458, 2005.[CrossRef][Web of Science][Medline]
  43. Strassmann G and Kambayashi T. Inhibition of experimental cancer cachexia by anti-cytokine and anti-cytokine-receptor therapy. Cytokines Mol Ther 1: 107–113, 1995.[Web of Science][Medline]
  44. Tidball JG. Inflammatory processes in muscle injury and repair. Am J Physiol Regul Integr Comp Physiol 288: R345–R353, 2005.[Abstract/Free Full Text]
  45. Tisdale MJ. Cachexia in cancer patients. Nat Rev Cancer 2: 862–871, 2002.[CrossRef][Web of Science][Medline]
  46. Tsujinaka T, Ebisui C, Fujita J, Kishibuchi M, Morimoto T, Ogawa A, Katsume A, Ohsugi Y, Kominami E, and Monden M. Muscle undergoes atrophy in association with increase of lysosomal cathepsin activity in interleukin-6 transgenic mouse. Biochem Biophys Res Commun 207: 168–174, 1995.[CrossRef][Web of Science][Medline]
  47. Tucker JM, Davis C, Kitchens ME, Bunni MA, Priest DG, Spencer HT, and Berger FG. Response to 5-fluorouracil chemotherapy is modified by dietary folic acid deficiency in Apc(Min/+) mice. Cancer Lett 187: 153–162, 2002.[CrossRef][Web of Science][Medline]
  48. Yan Z, Choi S, Liu X, Zhang M, Schageman JJ, Lee SY, Hart R, Lin L, Thurmond FA, and Williams RS. Highly coordinated gene regulation in mouse skeletal muscle regeneration. J Biol Chem 278: 8826–8836, 2003.[Abstract/Free Full Text]
  49. Zuijdgeest-van Leeuwen SD, van den Berg JW, Wattimena JL, van der Gaast A, Swart GR, Wilson JH, and Dagnelie PC. Lipolysis and lipid oxidation in weight-losing cancer patients and healthy subjects. Metabolism 49: 931–936, 2000.[CrossRef][Web of Science][Medline]



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