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(The FASEB Journal. 2005;19:1431-1440.)
© 2005 FASEB

Proton NMR spectroscopy shows lipids accumulate in skeletal muscle in response to burn trauma-induced apoptosis

Loukas G. Astrakas*,{dagger}, Igor Goljer{ddagger}, Shingo Yasuhara§, Katie E. Padfield*, Qunhao Zhang*, Suresh Gopalan*, Michael N. Mindrinos{cjs0822}{cjs0822}, George Dai{dagger}, Yong-Ming Yu*, J. A. Jeevendra Martyn§, Ronald G. Tompkins*, Laurence G. Rahme* and A. Aria Tzika*,{dagger},1

* Department of Surgery, Massachusetts General Hospital, Shriners Burns Institute and Harvard Medical School, Boston, Massachusetts, USA;
{dagger} Athinoula A. Martinos Center of Biomedical Imaging, Department of Radiology, Massachusetts General Hospital, Boston, Massachusetts, USA;
{ddagger} Varian NMR Systems, Columbia, Maryland, USA;
§ Departments of Anesthesiology and Critical Care, Massachusetts General Hospital, Shriners Burns Institute and Harvard Medical School, Massachusetts; and
{cjs0822}{cjs0822} Department of Biochemistry, Stanford University School of Medicine, Stanford, California, USA

1 Correspondence: NMR Surgical Laboratory, Department of Surgery, Massachusetts General Hospital, Harvard Medical School, 51 Blossom St., Room 261, Boston, MA 02114, USA. E-mail:atzika{at}partners.org


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Burn trauma triggers hypermetabolism and muscle wasting via increased cellular protein degradation and apoptosis. Proton nuclear magnetic resonance (1H NMR) spectroscopy can detect mobile lipids in vivo. To examine the local effects of burn in skeletal muscle, we performed in vivo 1H NMR on mice 3 days after burn trauma; and ex vivo, high-resolution, magic angle spinning 1H NMR on intact excised mouse muscle samples before and 1 and 3 days after burn. These samples were then analyzed for apoptotic nuclei using a terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL) assay. To confirm our NMR and cell biology results, we used transcriptome analysis to demonstrate that burn trauma alters the expression of genes involved in lipid metabolism and apoptosis. Our results demonstrate that burn injury results in a localized intramyocellular lipid accumulation, which in turn is accompanied by burn-induced apoptosis and mitochondrial dysfunction, as seen by the up-regulation of apoptotic genes and down-regulation of genes that encode lipid oxidation and the peroxisomal proliferator activator receptor {gamma} coactivator PGC-1ß. Moreover, the increased levels of bisallylic methylene fatty acyl protons (2.8 ppm) and vinyl protons (5.4 ppm), in conjunction with the TUNEL assay results, further suggest that burn trauma results in apoptosis. Together, our results provide new insight into the local physiological changes that occur in skeletal muscle after severe burn trauma.—Astrakas, L. G., Goljer, I., Yasuhara, S., Padfield, K. E., Zhang, Q., Gopalan, S., Mindrinos, M. N., Dai, G., Yu, Y.-M., Jeevendra Martyn, J. A., Tompkins, R. G., Rahme, L. G., Tzika, A. A. Proton NMR spectroscopy shows lipids accumulate in skeletal muscle in response to burn trauma induced apoptosis.


Key Words: magnetic resonance spectroscopy • high-resolution magic angle spinning skeletal muscle • burn trauma • mitochondria • intramyocellular lipids


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
A THOROUGH UNDERSTANDING of the physiology of burn trauma is important to the development of successful therapies for burn patients. Burn trauma in skeletal muscle has both local and systemic effects, as functionally debilitating changes are seen to occur at local and distant sites, especially when burn size exceeds 30% of total body surface area (1 , 2) . Burn patients exhibit an array of metabolic dysfunctions, including severe hypermetabolism, catabolism associated with accelerated gluconeogenesis, glucose oxidation and clearance, ineffective cycling of carbohydrate intermediates, accelerated lipolysis and futile fatty acid cycling, and increased amino acid oxidation and urea synthesis (3) . Furthermore, burn trauma results in proteolysis, muscle wasting, and prolonged loss of lean body mass, which are difficult to limit and treat using current nutritional therapies (4) . Muscle wasting also accompanies a range of systemic diseases including cancer, chronic uremia, AIDS, sepsis, diabetes mellitus, and hyperthyroidism. Although these dysfunctions likely trigger muscle wasting via distinct extracellular stimuli, the resulting physiological changes share several features (5) . For instance, the rapid loss of muscle protein mainly results from increased cell protein degradation (6) , especially of contractile proteins.

Apoptosis or programmed cell death is an evolutionarily conserved process that targets the removal of infected, malignant, or developmentally redundant cells, and may serve as an alternative mechanism that mediates the loss of parenchymal tissue, including muscle that accompanies trauma (7) . Muscle wasting associated with burn trauma may initially be triggered by the release of circulating cytokines that activate apoptosis. A prime cytokine pathway candidate for such activation is tumor necrosis factor-{alpha} (TNF-{alpha}), which appears to be elevated by burn trauma (8) , as well as in several other disease states, and might trigger apoptosis via the TNFR1 intracellular death domain and the caspase cascade (9) . TNF-{alpha} activation of apoptosis is likely further provoked by ceramide, a product of sphingolipid metabolism (10 , 11) .

Nuclear magnetic resonance (NMR) spectroscopy has been used to explore metabolic changes after burn trauma in liver tissue extracts (12 , 13) . The accumulation of mobile lipids detected by NMR may be associated with many cellular processes including proliferation (14) , inflammation (15) , apoptosis (16) , and necrosis (17) . Mobile lipid accumulation, presumably due to apoptosis, has been observed by 1H NMR during glioma gene therapy (18) . In vivo 1H NMR has detected the accumulation of intramyocellular lipids in the elderly (19) and in patients with type 2 diabetes (20) , which was attributed to impaired mitochondrial activity. Indeed, that the core apoptotic pathway and cellular energy metabolism are the two major determinants of cellular survival suggests an important interrelationship exists between apoptosis and mitochondrial function (21) . Based on these and previous results (22) , we hypothesize that mobile lipid accumulation accompanies severe burn trauma in muscle, and that this accumulation is the result of apoptotic myocytes, to suggest that mitochondrial dysfunction is involved in the burn response. Here we apply NMR, histopathology, and whole-genome expression analyses to a mouse burn trauma model (23) to test this hypothesis and further characterize the direct effects of burn. Our results provide novel insight into the local physiology of burn trauma and suggest a candidate target through which muscle wasting in burn patients might be limited.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals
Six-wk-old 20–25 g CD1 male mice (Charles River Laboratory, Boston, MA, USA) were housed in a temperature-controlled room with a 12 h light cycle and provided standard rodent feed and water ad libitum.

Burn trauma mouse model
Mice were injured using an established burn trauma model (24 , 25) approved by the Massachusetts General Hospital Institutional Animal Research Review Board Committee. Mice were anesthetized by intraperitoneal injection of 40 mg/kg pentobarbital sodium (Abbott Laboratories, Chicago, IL, USA) and randomized into burn or control groups. An area of the left leg corresponding to 3–5% of the total body surface area was shaved, and the burn group mice were subjected to scald burn trauma of the exposed skin by immersion for 4 s in 90°C water. Buprenophine (0.05–0.1 mg/kg) was subcutaneously provided 8–12 hourly as analgesia. This study focuses on the effects of local burn vs. those of systemic burn, which are limited to thermal injury burn in excess of 30% total body surface area. To this end, we chose limb muscle tissue from unburned mice for the control vs. the unburned contralateral muscle from the burned mice. This eliminates the possibility that the unburned contralateral control muscle has undergone potential systemic effects that might alter muscle physiology and gene expression.

In vivo 1H NMR spectroscopy
An initial group of six mice was studied with in vivo 1H NMR spectroscopy 1 and 3 days after burn trauma, after 24 h fasting. On the day of the NMR experiment, mice were placed in a customized restraining tube to limit free movement of the secured leg to be studied. The mice were then anesthetized using a nose cone and vaporizer with a halothane/oxygen mixture (5% for induction, 1% for maintenance). The rectal body temperature was maintained at 37 ± 1°C using heated water blankets. All in vivo 1H NMR experiments were performed with a 2 cm diameter circular surface coil placed above the area of interest in an horizontal bore magnet (proton frequency at 400 MHz, 21 cm diameter, Magnex Scientific, Oxfordshire, UK) using a Bruker Avance console. Magnetic resonance imaging (MRI) was performed using a 3-dimensional rapid acquisition with relaxation enhancement (RARE) fast spin-echo sequence (26) with typical acquisition parameters of (TR=2 s, TE=32 ms, NEX=1, 10 slices, slice thickness=1 mm). Then 1H NMR spectra from the left hind limb burned muscle and the corresponding right hind limb nonburned muscle were acquired, from burned and sham control mice, using a localized stimulated echo acquisition mode (STEAM) (27) sequence TE = 16 ms, TR = 2 s, 40 averages, 4KHz spectral width, 2048 points. The XWIN NMR software package (Paravision X, Bruker) was used for peak quantitation. Free induction decays were zero filled to 8K points and apodized with exponential multiplication (2 Hz) before Fourier transformation. The spectra were then manually phased and corrected for baseline broad features. The Levenberg-Marquardt algorithm was used to least-squares fit a model of mixed Gaussian/Lorentzian functions to the data. Student’s t tests were used to compare the peak values between control, 1 day, and 3 day burned animals.

Ex vivo magic angle spinning (MAS) 1H NMR spectroscopy
Twelve burned mice were analyzed by MAS 1H NMR spectroscopy: 1 and 3 days postburn the mice were killed; and their skeletal muscle tissue underlying the hind limb burn site was harvested, immediately frozen in liquid nitrogen and stored at –80°C. Limb gastronemius muscle tissue excised from six unburned animals at the same time points as above served as controls. All samples were analyzed by MAS 1H NMR in a Varian 14.1 Tesla spectrometer at 4°C and 3190 Hz spinning speed. The Carr-Purcell-Meiboom-Gill (CPMG) (28) spin-echo pulse sequence [90°-({tau}-180°-{tau})n-acquisition] with a delay ({tau}) of 270 µs was used to measure spin echo MAS 1H NMR spectra on all samples. Typically, 256 transients were collected into 32 K data points; the total spin-spin relaxation delay after the 90° pulse was kept ~11.928 ms and the number of 180 degree cycles was n = 2. For assignment purposes, 1H-13C gradient heteronuclear single quantum coherence (HSQC) (29) NMR spectra were measured under MAS conditions, with typical parameters: 1H spectral width 7 ppm; 13C spectral width 150 ppm; number of transients 16; number of increments 400 complex in t1 dimension; presaturation of water resonance, in combination with gradient selection, was used to suppress the water signal; and spectra were acquired using the hypercomplex states method in the t1 dimension. Spectral processing was performed using linear prediction in t1 up to 1024 points with matched Gaussian apodization in both dimensions. The VNMR 6.1 software package was used for peak quantitation. Student’s t tests were used to compare the peak values between control and 1 and 3 day burned animals.

Cellular studies/histopathology
In situ terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL) was used to detect apoptotic nuclei (30) according to the manufacturer (Oncor, Gaithersburg, MD, USA). The tissue samples used above for MAS 1H NMR were cryosectioned into 10 µm thick slices, fixed in 4% paraformaldehyde for 10 min at room temperature, washed with PBS, postfixed in ethanol-acetic acid (2:1) for 5 min at –20°C, and incubated with terminal deoxynucleotide transferase. The subsequent fragmented DNA was visualized with fluoresceine-conjugated antibody, and nuclei and muscle were counterstained with 4',6-diamidino-2-phenylindole, anti-caveolin-3 antibody, and Texas Red-conjugated secondary antibody, respectively.

RNA extraction, transcriptome studies, and analysis
Skeletal muscle from the control and burn hind limbs 1 and 3 days post-trauma was excised and immediately immersed in 1 ml Trizol (Invitrogen, Carlsbad, CA, USA), homogenized for 60 s using a Brinkman Polytron 3000, and chloroform extracted. RNA was isolated by isopropanol precipitation and centrifugation at 12000 g for 10 min; purified using the RNeasy Kit (Qiagen, Germantown, MD, USA); quantified by UV absorbance at 260 and 280 nm; and stored at –80°C prior to microarray analysis. Using standard Affymetrix protocols (Affymetrix Inc., Santa Clara, CA, USA), biotinylated cRNA was generated from 10 µg of total cRNA, and hybridized onto MOE430A oligonucleotide slide arrays, which were stained, washed, and scanned.

For genomic data analysis, the image data files of the RNA probes from burn/trauma and control animals at 1 and 3 days were converted to cell intensity files (.CEL files) with the Microarray suite 5.0 (MAS, Affymetrix, CA, USA). The data was scaled to a target intensity of 500, and all possible pairwise array comparisons of the replicates were performed for each time point (i.e., four combinations consisting of two arrays from each time point vs. the two arrays from normal mouse), using a change call algorithm of MAS 5.0. Genes with change calls of the same direction in three of four comparisons, and a signal value difference > 100 in each comparison, with the non-zero signal log ratio and one of the two samples being compared not called "absent," were scored as differentially modulated. The ratio of differential expression is the average of all four inverse log2 values of the signal log ratio. The ratio for genes with decreased expression compared with control mice are expressed as negative values. An additional constraint of a minimum ratio of 1.65 was applied to set the known false positives at 5% [based on the ratios of 100 genes determined to be invariant in most conditions tested (Affymetrix, Santa Clara, CA, USA)] when a gene was identified as differentially expressed at just one time point.

Statistically significant sets of functionally related genes were selected from a set of 1136 genes identified to be differentially expressed in the mouse hind limb burn model vs. unburned control mice (31) . These subsets were identified and compiled from experimental evidence or annotations of apoptosis, mitochondrial or fatty acid oxidation using annotation of the MOE430A chip [Affymetrix, retrieved December, 2003; and GeneSpider function, [Gene Spring, Silicon Genetics, CA, USA), compiled from GenBank, Locuslink and Unigene (32) ]. The overrepresentation statistics were calculated as hypergeometric probablility using all genes selected in each experiment having Gene Ontology annotation for a biological process (33) , using information compiled for the MOE430A chip [Affymetrix; and from Kyoto Encyclopedia of Genes and Genomes pathways (34) (retrieved March, 2004)]. P values were calculated using the R statistical package, version 1.7.1 for Windows (35) . Functional categories that did not have at least three genes at one time point and overlapping categories were deleted.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
In vivo and ex vivo 1H NMR detect muscle lipid accumulation after burn trauma
The in vivo and ex vivo NMR spectroscopy results presented in Fig. 1 and Fig. 2 and Table 1 demonstrate a consistent increase in most lipid peaks in the burned vs. nonburned skeletal muscle; while metabolites, such as choline-containing compounds, total creatine, taurine, glucose, and glycogen do not significantly change in the muscle in response to burn trauma.



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  		Figure 1. In vivo 1HNMR spectra and image of mouse skeletal muscle after 1 day burn trauma or sham treatment. A) In vivo 1HNMR spectra of a burned (green line) and sham-treated (purple line) hind limb. Spectra are normalized to the total creatine peak at 3.02 ppm. IMCL, intramyocellular (curved arrow); B) T1 weighted images (using RARE, TR=2 s, TE=32 ms, NEX=1, 10 slices, slice thickness=1 mm) showing the location of the voxels of interest (VOI) (squares) in control and burned hind limbs. Dotted circles in the image outline skeletal muscle regions in burned (right) and contralateral (left) hind limbs of a typical mouse after experimental burn trauma. VOI correspond to the location of MR spectroscopy data acquisition; green square = burned, purple square = nonburned. In agreement with NMR spectroscopy (A), MR imaging exhibits differences, albeit nonspecific, in signal intensities between burned and control or contralateral hind limbs.



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  		Figure 2. Ex vivo MAS 1H NMR spectroscopy of skeletal muscle biopsies after burn trauma. Lipid resonances (gray shaded peaks) increase over time after burn. Key: 1, CH3–(CH2)n lipids; 2, (CH2)n lipids; 3, CH2–CH2–CO lipids; 4 CH=CH–CH2–CH2 lipids; 5 CH2–CH2–CO lipids; 6, =CH–CH2–CH= lipids; 7, Total creatine (creatine and phosphocreatine); 8, choline-containing compounds; 9, taurine; 10, glucose; 11, glycogen; 12 residual water; 13 CH–O-CO- lipids; 14 –CH=CH- lipids.


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  		Table 1. 1H NMR lipid analysis of mouse skeletal muscle at day 1 and day 3 postburna

Figure 1A presents representative in vivo 1H MRS results from burned and nonburned leg muscles of the same mouse, and Fig. 1B shows axial images at the level of the hind limb and the 1H MRS data acquisition sites. The spectra have been scaled to the phosphocreatine and creatine peak (3.02 ppm). Resonance signals are due to residual water (4.7–4.8 ppm), terminal methyl (0.8–1.0 ppm), acyl chain methylene (1.1–1.5 ppm), {alpha}- and ß-methylene (2.0–2.5 ppm) and olefinic protons (5.4 ppm) of lipids, N-methyl protons of phosphocreatine and creatine (3.0 ppm) and N-trimethyl protons of betaines (3.2 ppm) comprising taurine and choline-containing compounds. The black arrow indicates the lipid peak at 1.4 ppm, where the greatest difference between the two experimental paradigms occurs. A similar increase after burn trauma is also seen for lipids at 2.8 ppm, which contains the bis-allylic methylene group, whereas the 1.1 ppm and 0.9 ppm lipid peaks decrease with trauma. Difficulties inherent to short echo-time (TE) localized in vivo spectroscopy could preclude reliable estimation of the response of specific lipid peaks and produce conflicting results between our in vivo and ex vivo NMR data. Such difficulties include incomplete water suppression, broad baseline features, poor shimming, and physiological noise as well as potential artifacts due to subcutaneous fat contamination in the absence of outer volume suppression. The Fig. 1B image demonstrates signal intensity differences possibly attributable to nonspecific lipid changes that occur in the burned vs. nonburned sites. Quantitative results from the higher resolution ex vivo MAS 1H MRS measurements verify the in vivo MRS results (Table 1) .

The Fig. 2 stack plot presents typical examples of proton MAS 1H NMR CPMG spectra from skeletal muscle. This plot contains spectra of control and burn animals killed 1 and 3 days post-trauma, with the spectra scaled to the phosphocreatine and creatine 3.02 ppm peak. Resonance signals are due to residual water (4.7–4.8 ppm); terminal methyl (0.8–1.0 ppm); acyl chain methylene (1.1–1.5 ppm); {alpha}- and ß-methylene (2.0–2.5 ppm) and olefinic protons (5.4 ppm) of lipids; N-methyl protons of phosphocreatine and creatine (3.0 ppm); and N-trimethyl protons of betaines (3.2 ppm), which correspond to taurine and choline-containing compounds. Particular lipid peaks are shaded to emphasize their increase over time after burn, with large differences between burned and sham-burned tissue primarily observed in the lipid peaks. The phospholipids at 3.2 ppm appear unaffected by trauma and remain stable, along with other peaks between 3 and 4 ppm. Quantitative results from these high-resolution ex vivo MAS 1H MRS measurements are shown in Table 1 . Furthermore, the 2-dimensional (2-D) heteronuclear single quantum coherence (HSQC) MAS 1H–13C NMR results, presented in Fig. 4 , confirm the 1-dimensional 1H spectral assignments via 13C correlations and literature values (36 , 37) .



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  		Figure 4. Heteronuclear single quantum coherence spectrum from the skeletal muscle of a control mouse. Key: 1, CH=CH–CH2–CH2 lipids; 2, CH2–CH2–CO lipids; 3, CH=CH–CH2–CH2 lipids; 4, Creatine and phosphocreatine; 5, Hypo-Taurine; 6, Taurine; 7, choline-containing compounds; 8, Creatine; 9, C6 {alpha}/ß glucose; 10, C6H glycogen; 11, C2H glycogen; 12, CH=CH–CH2– or CH=CH– lipids.

In situ TUNEL demonstrates apoptosis in response to burn trauma
Figure 3 presents a representative in situ TUNEL assay that demonstrates that apoptosis occurs postburn. Normal nuclei stain blue and apoptotic nuclei green. The observed apoptosis is not due to cells infiltrating from outside, as apoptotic nuclei occur within the area stained red by caveolin-3, a skeletal muscle protein marker. The number of apoptotic nuclei is higher in samples at day 3 vs. day 1 postburn, in agreement with previous results (22) . Indeed, on day 3 the TUNEL-positive nuclei in muscle samples of burned animals vs. the normal controls were 79.6 (mean) ± 14.1 (SE) and 0.86 (mean) ± 0.37 (SE), respectively, which is significantly different [P<0.01 by Mann-Whitney Rank-Sum Test (n=6)].



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  		Figure 3. In situ terminal deoxyribonucleotidyl transferase nick end labeling (TUNEL) assay of hind limb skeletal muscle from control and burned mice. Nuclei are stained blue with 4',6-diamidino-2-phenylindole (DAPI), and apoptotic nuclei are stained green with fluoresceine-conjugated antibody (TUNEL FITC). Apoptosis was present after burn day 1. On day 3, the apoptosis, evidenced as green staining, appeared in muscle and was most prominent at 3 days after burn as compared with day 1. Anti-caveolin-3 staining (Cav3-TexRed) shows that muscle structure was unaltered at 1 day but is gradually disorganized by 3 days. Green and red images are superimposed (TUNEL+Cav3) to show that most of the apoptotic nuclei are within muscle cells and not outside due to infiltrating cells.

Two-dimensional NMR confirms the 1-dimensional NMR spectral assignments
Two-dimensional heteronuclear single quantum coherence (HSQC) MAS 1H-13C NMR, presented in Fig. 4 , confirms the 1H resonance assignments via 13C correlations and literature values (36 , 37) .

Microarray analysis suggests burn trauma results in apoptosis and mitochondrial dysfunction
We seek to identify all the skeletal muscle genes whose expression is altered in response to burn trauma. To this end, we have carried out whole-genome expression analysis of the mouse hind limb burn model and identified 1136 genes that are differentially expressed vs. unburned control mice (31) . In concert with our NMR and TUNEL data, we identified a subset of the differentially expressed genes that have functional annotations of apoptosis and fatty acid metabolism. Of the initial 1136 genes, 19 act in apoptosis, with three encoding mitochondrial-dependent apoptosis functions; 10 are involved in fatty acid oxidation.

Differential expression of apoptosis-related genes
The microarray data presented in Table 2 show that 19 genes, identified to function in apoptosis via annotation by the Gene Ontology Consortium and Ingenuity (Ingenuity Pathway Analysis), are differentially expressed and that 14 of these genes are up-regulated and 6 are down-regulated for at least one time point after burn injury vs. controls. For example, the TNF receptor superfamily member 1a gene shows increased expression at 1 day and 3 days postburn. This gene encodes a major TNF-{alpha} receptor that activates NF-{kappa}B, mediates apoptosis, and regulates inflammation (38) . Genes encoding the Bcl2-associated X protein (BAX) and baculoviral IAP repeat-containing 2 (cIAP), which function downstream of the TNF-{alpha} receptor in apoptosis signaling, exhibit increased expression at 3 days. Several other genes are up-regulated, including the death-associated protein kinase 1, growth arrest, and DNA damage-inducible 45 gamma genes; and the CCAAT/enhancer binding protein (C/EBP) gene, whose product regulates immune and inflammatory response genes, and binds acute-phase and cytokine genes. Genes primarily up-regulated at day 1 encode the anti-apoptosis heat shock proteins 1B and 70 kDa (39) , and immediate early response 3. Mitochondrial-dependent apoptosis genes that encode cytochrome c, caspase 7, caspase-regulated phosphodiesterase 4A, and DNA fragmentation factor (ICAD) show decreased expression.


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  		Table 2. Differential expression of apoptosis genes at day 1 and/or day 3 postburn traumaa

Fatty acid oxidation
Ten fatty acid oxidation genes are differentially expressed after burn trauma and exhibit reduced expression (Table 3 ), nine of which are down-regulated 3 days post-trauma. These genes encode the lipoprotein lipase and adipocyte complement-related proteins, which respectively increase the non-esterified fatty acids cytosolic pool and functions in fatty acid ß-oxidation, as well as dodecenoyl-CoA delta isomerase, acetyl-CoA acyltransferase 2, and carnitine palmitoyltransferase 1 (CPT1), the rate-controlling enzyme of the long-chain fatty acid ß-oxidation pathway in muscle mitochondria. Reduced CPT1 activity could contribute to muscle lipid accumulation and the lipid metabolism dysfunction associated with burn injury. The peroxisomal proliferator activator receptor {gamma} coactivator PGC-1ß gene is down-regulated at both time points, in agreement with the general down-regulation of fatty acid oxidation genes. PGC-1ß belongs to the PGC-1 transcriptional regulator family that is responsible for mitochondrial biogenesis and fatty acid oxidation (40) .


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  		Table 3. Differential expression of fatty acid oxidation genes at day 1 and/or day 3 postburna


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Energy expenditure in severe burn trauma increases ~57% due to major ATP-consuming reactions (41) . Here, we employ ex vivo and in vivo NMR, histopathology, and genomics studies to examine the metabolic and gene expression changes that occur at local sites of burn injury. Our data demonstrate that burn trauma leads to increased accumulation of intramyocellular lipids associated with apoptosis and defects in specific mitochondrial pathways. These results provide further insight into the biochemistry and physiology of local burn trauma.

While in vitro NMR spectroscopy has been used to explore metabolic changes after burn trauma in liver tissue extracts (12 , 13) and in bacterial and cell cultures (42 , 43) , its limited sensitivity has prohibited its in vivo application. Magic angle spinning (MAS), an NMR technique used for solids, can produce high-resolution spectra for biological samples. Compared with conventional high resolution in vitro NMR of tissue extracts, ex vivo MAS NMR of intact tissues can provide superior biochemical information, with simplified sample preparation and a better approximation of the in vivo state. This technique is increasingly being used to investigate overt cellular diseases (44 45 46 47) . Here we apply MAS 1H NMR to intact muscle samples from animals subjected to burn-trauma to identify at high-resolution concomitant metabolic and molecular aberrations associated with apoptosis. These changes might also reflect mitochondrial dysfunction, as some genes whose expression is down-regulated in response to burn encode mitochondrial-dependent apoptosis functions.

A principal finding of our experiments is that mobile lipids accumulate in muscle tissue in response to burn trauma. Although the source of these accumulated lipids is beyond the scope of this study, it has been shown that extramyocellular lipids (EMCL), intramyocellular lipids (IMCL), and triglycerides can all contribute to cellular lipid peaks (48 49 50) . Indeed, EMCL and IMCL can be measured by in vivo MRS due to differences in bulk magnetic susceptibility and geometric arrangements (48 49 50) . IMCL probably serve as an energy substrate for oxidative metabolism (51) , and can be mobilized and used with turnover times of several hours (52) . Furthermore, the lipid peak at 1.4 ppm in Fig. 1 has been attributed to methylene protons of intramyocellular triglyceride acyl chains, primarily due to IMCL (49) , to suggest that the increase in NMR-visible lipids at 1.4 ppm after burn-trauma is primarily due to increased quantifiable IMCL. This conclusion is further supported by the results of human studies (19 , 20) . Conversely, EMCL are relatively metabolically inert and serve as a long-term energy storage depot with slow turnover. EMCLs are therefore unlikely to correspond to the lipids observed here that quickly accumulate postburn trauma. We note that the poor sensitivity of in vivo NMR, as well as subcutaneous contamination of the experimental tissue, might explain differences between the in vivo and ex vivo results. The elevated methyl and methylene proton resonances may correspond to cytoplasmic mobile protein moieties (53 , 54) . We also note that we acquire the ex vivo NMR spectra from untreated muscle samples and heavily T2 weighted MAS CPMG sequences, and thus, as expected, these spectra lack the detailed pattern characteristic of spectra obtained postpurification using liquid NMR methods.

Apoptosis, necrosis, and inflammation likely perturb muscle lipid content after burn trauma to explain the observed NMR changes. Indeed, lipids are associated with these cellular processes (14 15 16 17) ; and the Fig. 3 TUNEL assay data provide the first demonstration that burn trauma triggered apoptosis and the increased lipid profile, seen in Fig. 2 , are linked. The increase of bisallylic methylene fatty acyl protons at 2.8 ppm agrees with data in Hakumaki et al. (18) , which suggested these protons correspond to polyunsaturated fatty acids (PUFAs) and PUFA accumulation that follows apoptosis. In addition, our data demonstrate vinyl proton accumulation at 5.4 ppm, including protons from ceramide and possibly other sphingolipids, such as sphingosine and other monounsaturated fatty acids. These enhanced signals suggest the sphingolipid pathway contributes to burn-trauma mediated apoptosis. Indeed, sphingolipids are involved in apoptosis and burn injury (22 , 55) . Although the upstream signaling of apoptosis remains uncertain, sphingolipids, and ceramide in particular, have been implicated as second messengers for stress stimuli, including TNF-{alpha} (56) , Fas ligand (57) , ionizing radiation (58) , heat shock (55) , ultraviolet light (59) , and oxidative stress with SAPK activation and mitochondrial translocation (60) . Peaks in the ceramide resonance region have been detected in brain gliomas (61) . Although in vivo NMR does not readily detect ceramide or sphingolipids due to their limited biological concentrations, MAS NMR can detect their monounsaturated fatty acid precursors. Absolute identification of the ceramide-related contribution to the NMR peaks, and of its metabolic involvement in burn trauma, will require additional biochemical approaches. Nevertheless, the NMR spectra, in conjunction with the TUNEL assay data, further suggest that burn trauma mediates apoptosis and induces the sphingolipid pathway, including ceramide, an intracellular apoptosis signal.

Our transcriptome studies further demonstrate that apoptotic gene expression is altered in response to burn trauma (Table 2) . As these genes encode proteins having pro- or anti-apoptotic functions, their exact regulation remains unclear. These data may reflect a heterogeneous mRNA population due to a direct effect of burn trauma on the underlying superficial layers of the hind limb muscle and the asynchronous state of apoptotic muscle cells. Nonetheless, the data demonstrate that apoptosis pathways are activated in the muscle postburn to further demonstrate a link between lipid accumulation and apoptosis. That mitochondrial-dependent apoptosis genes exhibit decreased expression suggests mitochondrial impairment might also play a role in the burn response. Indeed, our recent data (31) show that mitochondrial dysfunction accompanies burn pathogenesis in skeletal muscle with alterations in mitochondrial-directed energy expenditure reactions, including down-regulation of mitochondrial oxidative phosphorylation.

IMCL accumulation suggests that enzymes responsible for lipid oxidation are down-regulated, as verified by the genomics data. Significantly, this down-regulation identifies a specific defect responsible for IMCL accumulation that could assist in the interpretation and treatment of burn patients. The observed reduced expression of PGC-1ß, a transcriptional regulator of mitochondrial biogenesis and fat oxidation genes (40 , 62 , 63) , further demonstrates that fat oxidation, a process largely localized to the mitochondria, is reduced after severe local burn trauma. PGC-1 down-regulation has also been linked to type 2 diabetes (64 , 65) , and PGC-1 regulates the expression of multiple nuclear respiratory factor-1 (NRF-1) -dependent genes, such as cytochrome c oxidase subunits and ATP synthases (66) . Indeed, these key enzymes in oxidative metabolism and mitochondrial function show progressive down-regulation 3 days postburn (data not shown).

Increased IMCL is associated with insulin resistance, a major metabolic dysfunction of burn trauma (67 , 68) . Previous measurements of muscle triglyceride content by biopsy (69) and IMCL content by 1H NMR spectroscopy (49 , 70 , 71) show a strong relationship between intramuscular fat content and insulin resistance in muscle. While increased fatty acid delivery from lipolysis could also produce the observed IMCL increase, free fatty acid concentrations are highly variable in burn patients (72) . Impaired lipoprotein and polyunsaturated fatty acid metabolism occurs in the early postburn period (73) , suggesting their involvement in subsequent healing and immune function. Our genomic data alternatively suggest that the increased IMCL could be the result of decreased mitochondrial oxidative capacity, in agreement with gene expression data in human diabetes (64) . It has been reported that increased IMCL is associated with insulin resistance in type 2 diabetes (20) , suggesting reduced mitochondrial oxidation and phosphorylation.

In conclusion, NMR, cell biology, and transcriptome results together provide strong evidence that the altered NMR visible lipid profile is related to apoptosis triggered by burn trauma. The results also suggest that this trauma leads to mitochondrial dysfunction, which could contribute to the increase in energy expenditure seen in burn patients. Lipid accumulation may reflect increased inflammation and apoptosis rather than providing a direct measure of mitochondrial dysfunction. Since apoptosis and cellular energy metabolism are the two major determinants of cell survival, NMR visible lipids may serve as biomarkers to monitor therapies of muscle wasting after burn trauma, as well as other disease states. To this end, the suppression of certain lipids such as ceramide and/or other sphingolipids may present a candidate strategy to limit muscle wasting after burn trauma.


   ACKNOWLEDGMENTS
 
We thank Dr. Scott Stachel for comments on the manuscript and editing. This work was supported in part by National Institute Institutes of Health (NIH) Center Grant P50GM021700, NIH Glue Grant U54GM062119, and Shriners Grant 8590 (to L.G.R.). Q.Z. is a Shriners Research search Fellow.

Received for publication June 17, 2004. Accepted for publication May 3, 2005.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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M. G. Cree and R. R. Wolfe
Postburn trauma insulin resistance and fat metabolism
Am J Physiol Endocrinol Metab, January 1, 2008; 294(1): E1 - E9.
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