FASEB J.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Published as doi: 10.1096/fj.07-8527rev.
 (The FASEB Journal. 2007;21:3431-3441.)
© 2007 FASEB
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
fj.07-8527revv1
fj.07-8527revv2
21/13/3431    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pieter de Lange
Right arrow Articles by Lanni, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pieter de Lange,
Right arrow Articles by Lanni, A.

Fuel economy in food-deprived skeletal muscle: signaling pathways and regulatory mechanisms

Pieter de Lange*, Maria Moreno{dagger}, Elena Silvestri{dagger}, Assunta Lombardi{ddagger}, Fernando Goglia{dagger} and Antonia Lanni*,1

* Dipartimento di Scienze della Vita, Seconda Università degli Studi di Napoli, Caserta, Italy;

{dagger} Dipartimento di Scienze Biologiche ed Ambientali, Università degli Studi del Sannio, Benevento, Italy; and

{ddagger} Dipartimento delle Scienze Biologiche, Sezione Fisiologia ed Igiene, Università degli Studi di Napoli "Federico II," Napoli, Italy

1Correspondence: Dipartimento di Scienze della Vita, Seconda Università degli Studi di Napoli, Via Vivaldi 43, 81100 Caserta, Italy. E-mail: antonia.lanni{at}unina2.it


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 
Energy deprivation poses a tremendous challenge to skeletal muscle. Glucose (ATP) depletion causes muscle fibers to undergo rapid adaptive changes toward the use of fatty acids (instead of glucose) as fuel. Physiological situations involving energy deprivation in skeletal muscle include exercise and fasting. A vast body of evidence is available on the signaling pathways that lead to structural/metabolic changes in muscle during exercise and endurance training. In contrast, only recently has a systematic, overall picture been obtained of the signaling processes (and their kinetics and sequential order) that lead to adaptations of the muscle to the fasting state. It has become clear that the reaction of the organism to food restraint or deprivation involves a rapid signaling process causing skeletal muscles, which generally use glucose as their predominant fuel, to switch to the use of fat as fuel. Efficient sensing of glucose depletion in skeletal muscle guarantees maintained activity in those tissues that rely entirely on glucose (such as the brain). To metabolize fatty acids, skeletal muscle needs to activate complex transcription, translation, and phosphorylation pathways. Only recently has it become clear that these pathways are interrelated and tightly regulated in a rapid, transient manner. Food deprivation may trigger these responses with a timing/intensity that differs among animal species and that may depend on their individual ability to induce structural/metabolic changes that serve to safeguard whole-body energy homeostasis in the longer term. The increased cellular AMP/ATP ratio induced by food deprivation, which results in activation of AMP-activated protein kinase (AMPK), initiates a rapid signaling process, resulting in the recruitment of factors mediating the structural/metabolic shift in skeletal muscle toward this change in fuel usage. These factors include peroxisome proliferator-activated receptor (PPAR){gamma} coactivator-1{alpha} (PGC-1{alpha}), PPAR{delta}, and their target genes, which are involved in the formation of oxidative muscle fibers, mitochondrial biogenesis, oxidative phosphorylation, and fatty acid oxidation. Fatty acids, besides being the fuel for mitochondrial oxidation, have been identified as important signaling molecules regulating the transcription and/or activity of the genes or gene products involved in fatty acid metabolism during food deprivation. It is thus becoming increasingly clear that fatty acids determine the economy of their own usage. We discuss the order of events from the onset of food deprivation and their importance.—de Lange, P., Moreno, M., Silvestri, E., Lombardi, A., Goglia, F., Lanni, A. Fuel economy in food-deprived skeletal muscle: signaling pathways and regulatory mechanisms.


Key Words: AMP-activated protein kinase • myosin heavy chain Ib • peroxisome proliferator-activated receptor {gamma} coactivator-1{alpha} • peroxisome proliferator-activated receptor {delta} • fatty acids • time course


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 
SKELETAL MUSCLE IS QUANTITATIVELY THE LARGEST ORGAN in the body, contributing 30–40% of the resting metabolic rate in adults. It is a major site for the oxidation of fatty acids and glucose (it accounts for ~80% of insulin-stimulated glucose uptake), and it exhibits a remarkable flexibility in its usage of fuel in response to energy deprivation. One notable aspect of skeletal muscle plasticity is the specificity with which it adapts to a given stimulus with structural, biochemical, and functional modifications (1) . This review deals with the timing of events in the signaling pathways involved in the response shown by skeletal muscle to food deprivation, taking into account data from other models of energy deprivation, transgenic models, and models involving hormonal/pharmacological stimuli, and it highlights the species differences found so far. Four major fiber types are recognized in skeletal muscle (2) . Type I fibers are slow twitch, showing a high capacity to synthesize ATP and that possesses many mitochondria and uses fatty acids over glucose as fuel (1 , 3) . Type II A, IIX, and IIB fibers have increasing capacities to utilize glucose and are increasingly fast twitch with reduced mitochondrial content (2) . Overall gene-expression analysis, including gene-array analysis, has revealed that the alterations occurring in skeletal muscle mRNA levels during energy deprivation are consistent with a suppression of anabolic pathways and an activation of catabolic pathways involving the uptake of glucose and fatty acid oxidation, events that together act to restore intracellular ATP levels (4 5 6 7) . The metabolic alterations that occur during food deprivation, leading to an increased flux of free fatty acids and changes in the AMP/ATP ratio, have evolved as major components of the phenotypic active stimuli in skeletal muscle, leading to fiber shifts. One factor considered to be central to ATP maintenance is AMP-activated protein kinase (AMPK; ref. 8 ), which is activated in response to elevated intracellular AMP levels. Muscle ATP levels can be restored by the administration of 5'-amino-4-imidazolecarboxamide (AICAR), which upon monophosphorylation is converted to the AMP analog zeatin riboside-S-monophosphate (ZMP) (9) , resulting in activation of AMPK in skeletal muscle (10 11 12) . This leads to a shift from type IIB fibers toward the more oxidative type IIX fibers (11) , without a shift toward type I fibers (11 , 12) . Such a fiber shift toward the type I phenotype is, however, triggered effectively by food deprivation. Indeed, within 6 h, food deprivation results in dramatic increases in the AMP/ATP ratio and AMPK phosphorylation (13) and elicits an increase in myosin heavy chain Ib (MHCIb) expression (13) , reflected by an accumulation of type I fibers in the longer term (13) . In accordance with this, it has been reported that low-frequency stimulation of muscle provokes an up-regulation of MHCIb in the rat (14) , albeit less dramatic than that provoked by food deprivation, and exercise training has been shown to induce differentiation of muscle fibers toward a fatigue-resistant, oxidative phenotype (3) . Fiber-type switching involves signaling mediated by 1) the key transcription factors peroxisome proliferator-activated receptor (PPAR){gamma} coactivator-1{alpha} (PGC-1{alpha}) and PPAR{delta}, 2) the energy-sensing and fuel-providing role of AMPK, 3) nitric oxide (NO), and 4) mitochondrial fatty acid (FFA) oxidation. Here, possible connections among these pathways in the processes after food deprivation will be highlighted. It should be noted that this review mainly focuses on the temporal aspects of the actions of the various factors discussed.


   FACTORS INVOLVED IN STRUCTURAL AND METABOLIC CHANGES IN SKELETAL MUSCLE UPON FOOD DEPRIVATION
TOP
 ABSTRACT
 INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 
PGC-1{alpha} is a signal-sensing coactivator, and in recent years, it has become clear that transcriptional coactivator proteins, apart from being crucial to the regulation of gene expression in response to cellular signals, can actually be the primary targets for hormonal control and signal-transduction pathways (for review, see ref. 15 ), also during low-energy states (see following section). These signal-controlled transcription factors include the PPAR{gamma} coactivator-1 (PGC-1) coactivators, which, through interaction with target transcription factors and other coactivators, induce powerful transcriptional activity (16 17 18 19) . One example of such PGC-1 coactivators is PGC-1{alpha}, which coactivates nuclear respiratory factors 1 and 2 (NRF-1 and NRF-2; ref. 20 ) and also nuclear hormone receptors, such as PPAR{alpha} (21) , PPAR{delta} (22) , estrogen-related receptor {alpha} (ERR{alpha}) (23 24 25 26 27) , and thyroid hormone receptor ß1 (TRß1) (28) . All of these transcription factors, their regulation, and their cellular targets have been discussed in greater detail in recent reviews (19 , 29 30 31 32) .

An important target tissue for PGC-1{alpha} is skeletal muscle (33) . As mentioned before, PGC-1{alpha} is a coactivator of PPAR{delta} (22) , and indeed a physical interaction between PGC-1{alpha} and PPAR{delta} has been demonstrated (by coimmunoprecipitation) in the nuclei of gastrocnemius muscles obtained from PPAR{delta} transgenic mice (22) . Experiments with transgenic mice carrying either PGC-1{alpha} or PPAR{delta} have revealed a surprising, clear role for both factors in inducing a muscle fiber-transition from glycolytic (type II) to oxidative (type I) (33 , 34) . Experiments on mice overexpressing a VP-16-activated form of PPAR{delta} have revealed that type I fiber formation can be directly stimulated without an induction of PGC-1{alpha} (34) , but in physiological situations an up-regulation of PGC-1{alpha} is likely to be involved in the early events leading to muscle-fiber shifts. Indeed, PGC-1{alpha}-null mice show clear muscle dysfunction (35) . In accordance with this, food deprivation causes a rapid, transient up-regulation of PGC-1{alpha} mRNA and nuclear protein and concomitantly induces a sharp increase in MHCIb mRNA and protein levels, indicative of the formation of type I fibers (13) . In fact, in the long term, this results in an accumulation of type I fibers in gastrocnemius muscle (13) . A positive correlation between PGC-1{alpha} and MHCIb expressions has also been made in the soleus muscle of rats stimulated with growth hormone (36) . Rapid up-regulation of PGC-1{alpha} in states of low energy has been shown to be achieved by various pathways involving AMPK (11 , 37 38 39) and NO (40) . Both pathways, and their possible interrelationship during food deprivation, will be discussed later in this review. An overview of the pathways leading to transcriptional changes during food deprivation is given in Fig. 1 .


Figure 1
View larger version (36K):
[in this window]
[in a new window]

  		Figure 1. Scheme depicting an in vivo short-term regulatory role for AMPK in transcriptional pathways in skeletal muscle after food deprivation. Rapidly increased AMP levels induce LKB1-mediated phosphorylation of AMPK, inhibited by an increase in intracellular fatty acids (FFA). Another trigger for AMPK-phosphorylation may be NO signaling. Once phosphorylated, AMPK stimulates 1) mitochondrial uptake of FFA for oxidation as an energy source, by phosphorylating ACC, thus indirectly activating the CPT system; and 2) transcription of PGC-1{alpha}, which in turn induces transcription of various factors, including PPAR{delta}. Increased nuclear accumulation of both PGC-1{alpha} and PPAR{delta} induces transcription of their target genes that are responsible for structural/metabolic changes manifested after food deprivation. White arrow pointing upward depicts a stimulatory effect. Triangles represent free fatty acid molecules. TF = transcription factors coactivated by PGC-1{alpha}; all other abbreviations are described in text.

With respect to PPAR{delta} and its targets, the high expression of PPAR{delta} in skeletal muscle (at 10- and 50-fold higher levels than PPAR{alpha} and PPAR{gamma}, respectively; see refs. 6 and 41 ), and its preferential expression in oxidative rather than glycolytic myofibers (34) , have led to the suggestion that this receptor may be involved in promoting the utilization of fatty acid as fuel. As mentioned above, studies on transgenic mice harboring a constitutively activated form of PPAR {delta} (VP16-PPAR{delta}) have clearly shown that this receptor plays a central role in the control of skeletal muscle lipid metabolism as well as in promoting the formation of type I, oxidative slow-twitch fibers (34) . A wild-type PPAR{delta} transgene (specifically overexpressed in skeletal muscle but not constitutively activated) has been shown to promote a net increase of fibers with an oxidative metabolic capability through an increment of total fiber number in soleus and tibialis anterior muscle, while in plantaris muscle the increase was more related to a shift from glycolytic to more oxidative fibers. Interestingly, the nonactivated PPAR{delta} transgene failed to induce the formation of type I slow in skeletal muscle (42) . In the latter case, despite its high expression, the actual activation of the transgene depends on the presence of natural ligands, such as fatty acids. These are present at fluctuating concentrations and may thus tightly regulate PPAR{delta} activity. These data raised a question as to whether, instead of overexpression, increasing the availability of ligand (natural or artificial) would induce fuel as well as fiber switching toward type I fibers. Recently, a fuel-switching role of PPAR{delta} has been demonstrated by incubating rat isolated skeletal muscle with the agonist GW50156 for 24 h. This led to an increased use of fatty acids over glucose, as reflected by increased fatty acid oxidation and reductions in glucose oxidation, glycogen synthesis, lactate release, and glucose transport. Interestingly, this switch was independent of insulin stimulation or fiber type (43) . In addition, prolonged treatment of mice with the same agonist not only increased the expression of genes involved in fatty acid oxidation, mitochondrial respiration, and oxidative metabolism (34 , 44) but also induced formation of type I fibers (34) .

Since situations of food shortage imply an increased use of fatty acids, it was anticipated that PPAR{delta} would be up-regulated and activated during fasting. However, the initial reports appearing in the literature initially did not seem to confirm this. Thus, a 24 h fasting period did not up-regulate PPAR{delta} in rat skeletal muscle (41) , and our group even reported reduced levels of PPAR{delta} (and PPA{alpha}) in gastrocnemius muscle from 48 h fasted rats (6) . Decreased expression of PPAR{delta} and PPAR{alpha} upon 48 h of fasting was recently reported in humans also (45) . In contrast, in the 24 h fasted mouse, skeletal muscle PPAR{delta} levels were found to be up-regulated (46) . These data indicate that single time point measurements may be misleading, due to possible transient modifications. In fact, time-course studies have shown that PPAR{delta} is indeed up-regulated in fasted rat skeletal muscle but within the first 6 h (13) . In addition, it has been observed that, like the up-regulation of PGC-1{alpha}, the up-regulation of PPAR{delta} during fasting is transient (13) . Rapid nuclear accumulation of both PGC-1{alpha} and PPAR{delta} upon food deprivation (13) and their physical interaction in the nucleus (22) are concomitant with increased fatty acid levels and increased expression of MHC Ib, which underlines the role of PPAR{delta} as a key regulator of fatty acid metabolism and muscle-fiber switching (in concert with its coactivator PGC-1{alpha}). Consequently, within 12 h of food deprivation, the PPAR-regulated genes mitochondrial thioesterase I (MTE I) (47) and carnitine palmitoyl transferase 1(CPT1), as well as uncoupling protein 3 (UCP3) (34 , 44) , were up-regulated simultaneously and so was the rate of mitochondrial fatty acid oxidation (13) .

The reported kinetics of PPAR{delta} up-regulation during the recovery period after exercise-imposed energy stress in humans are in line with those seen in the fasting rat: it has been shown that in humans, a single exhaustive bout of cycling increases PPAR{delta} mRNA and protein expression within 3 h after completion of the exercise (48 , 49) . This indicates that in general challenges involving immediate energy deprivation in skeletal muscle elicit rapid signaling through PPAR{delta}, thereby increasing oxidative capacity.


   MAINTENANCE OF SKELETAL MUSCLE ATP AND FUEL DURING FOOD DEPRIVATION
TOP
 ABSTRACT
 INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 
AMPK is an energy-sensing fuel provider. The need for an adequate response to metabolic stresses, such as fasting or exercise, was probably the evolutionary origin of a role for AMPK in single-celled eukaryotes (8) , and therefore this kinase is generally termed the "fuel provider" of the cell. Activation of AMPK through AMP is brought about as follows: muscle cells, like all eukaryotic cells, express the enzyme adenylate kinase, which catalyzes the reaction 2ADP {leftrightarrow} ATP +AMP and maintains it close to equilibrium at all times. In conditions involving ATP depletion (such as fasting), adenylate kinase activity increases so as to counteract the drop in ATP levels, which results in a rise in AMP levels. Thus, the AMP:ATP ratio is indicative of a compromised energy status (8) . It has been demonstrated, both in cultured cells (50) and in skeletal muscle in vivo (51) , that the upstream kinase that activates AMPK in response either to AMP-elevating treatments or to AMP-mimetic agents is LKB1. The LKB complex itself is not regulated by AMP and appears to be constitutively active (52 , 53) , with activation of the cascade being induced by the binding of AMP to AMPK, causing it to become a better substrate for LKB1 (50) . AMPK is known to be a heterotrimeric complex comprising a catalytic {alpha} subunit and regulatory ß and {gamma} subunits. Each subunit is encoded by multiple genes ({alpha}1, {alpha}2, ß1, ß2, {gamma}1, {gamma}2, and {gamma}3) yielding at least 12 heterotrimeric combinations, with splice variants further adding to the diversity. The {alpha}1 and {alpha}2 subunits are both phosphorylated and activated by LKB at position Thr-172 (50 51 52) . The {alpha}1 subunit is expressed in the oxidative/glycolytic type IIA and oxidative type I fibers, whereas the {alpha}2 subunit is expressed in all fiber types (54) . An obvious question that arises is whether both isoforms represent a redundancy or have specific activities.

In line with its central role, once activated, AMPK regulates PGC1-{alpha} expression in skeletal muscle, thereby boosting oxidative metabolism (11 , 37 38 39) . Indeed, physiological activation of AMPK, by food deprivation or exercise, is tightly linked to changes in the mRNA and protein expressions of PGC-1{alpha} and related genes. Recently, food deprivation has actually been shown to induce phosphorylation of AMPK and PGC-1{alpha} expression in skeletal muscle, with rapid kinetics (13) . Indeed, after a short fasting stimulus (6 h), skeletal muscle AMP levels rose and AMPK phosphorylation was increased, concomitant with an increase in PGC-1{alpha} expression (13) . Interestingly, the reported kinetics of AMPK activation during fasting are similar to those found during exercise: thus, 6 h of low-intensity swimming in rats resulted in increases in AMPK activation and PGC-1{alpha} mRNA expression (38) , and 30 min after the completion of a volontary treadmill run, AMPK activity increased in rat skeletal muscle (55) . A role has been demonstrated for AMPK in the fasting-induced up-regulation of the coordinated expression of genes controlling glucose and lipid metabolism in mice, since in AMPK {gamma} subunit-knockout mice starved for 16 h this up-regulation was abolished (56) . In knockout mice lacking the {alpha}2-, but not the {alpha}1-AMPK subunit, AMPK activation was markedly diminished during running. Nevertheless, exercise-induced activation of the investigated genes in mouse skeletal muscle was not impaired in {alpha}1- or {alpha}2-AMPK knockout muscles (57) . Although the authors of that paper felt the evidence did not favor an active role of AMPK in exercise-induced gene expression, in line with the question asked previously, a plausible hypothesis is that activation of the remaining {alpha}-subunit is sufficient to increase gene activation during exercise.

AMPK exerts its fuel-providing role by being activated in response to decreased fuel availability (glucose or fatty acids), and the fuel itself has been shown to play a critical role in the regulation of AMPK activity. Ex vivo incubation of the rat extensor digitorum longus muscle with high or low doses of glucose has revealed that AMPK activity was diminished at a raised glucose concentration (and vice versa), a result demonstrating the involvement of AMPK in the autoregulation of glucose uptake (58) . Furthermore, recent data demonstrated that experimental inhibition of oxidative phosphorylation by treatment with azide in Clone 9 rat liver cells (expressing GLUT 1), which leads to depletion of ATP levels and an increased demand for glucose uptake, correlate with an increased AMPK phosphorylation. Glucose uptake was shown to depend fully on AMPK-{alpha}1 in these experiments since inactivation of this subunit (through RNA interference) abolished glucose uptake in these conditions (59) . Clearly, AMPK activity needs to be tightly regulated to prevent excessive fuel usage, especially under circumstances of energy deprivation. In addition, in this case, the fuel itself plays a regulatory role. By performing time-course studies in food-deprived rats, we have recently demonstrated that the increased serum levels of FFA triggered by fasting exert a negative influence over AMPK phosphorylation in rat skeletal muscle (see Fig. 1 ; the results are schematically represented in Fig. 2 ). Shortly after the rapid, initial increase in AMPK phosphorylation, it was rapidly dephosphorylated once serum and intramuscular FFA levels began to rise (after ~12 h). In addition, in vivo nicotinic acid treatment, causing a rapid drop in serum FFA levels, led to rephosphorylation of AMPK. Throughout, skeletal muscle AMPK total protein levels did not change (13) . Additional proof that AMPK activity is regulated through feedback pathways associated with increased lipid metabolism has come from experiments on PGC-1{alpha}-deprived mice, which exhibit reduced fatty acid oxidation, with their livers developing steatosis on fasting (35) . In such mice, AMPK is constitutively activated in skeletal muscle, indicating an energy (fuel) deficit (60) . Given the link between AMPK activity and PGC-1{alpha} expression, the transient nature of the AMPK phosphorylation observed during fasting in gastrocnemius muscle of the food-deprived rat may explain the transient induction pattern of PGC1-{alpha} (with consequential transient coactivation of PPAR{delta}) and of the genes involved in lipid metabolism (13) .


Figure 2
View larger version (11K):
[in this window]
[in a new window]

  		Figure 2. Time-course data for transient induction of AMP levels, AMPK phophorylation, and subsequent rise in serum FFA levels, the last-named being responsible for subsequent dephosphorylation of AMPK.

In analogy to their role in the modulation of AMPK activity during fasting, there is accumulating evidence for a role for FFA in the inactivation of AMPK in obesity and obesity-related type 2 diabetes. Intriguingly, young insulin-resistant and obese Zucker diabetic fatty rats have been found to show defects in skeletal muscle AMPK phosphorylation and in PGC-1{alpha} protein expression levels, with endurance exercise on a treadmill partially correcting these abnormalities (61) . Another recent study (62) supports this idea, as rats fed a high-fat diet for 5 months had reduced skeletal muscle AMPK activity and reduced AMPK-{alpha} mRNA and protein expression levels, whereas pharmacological activation of AMPK by treatment with metformin increased both AMPK activity and AMPK-{alpha} mRNA and protein. The above-mentioned studies on rats have demonstrated a link between lipid accumulation and impaired AMPK activity in skeletal muscle, whereas initially human studies (63 , 64) have not provided evidence of any similar associations. However, very recently, time-course experiments with human subjects have revealed that exercise has a time- and intensity-dependent increasing effect on AMPK activity in muscle of lean human subjects, but obese and type 2 diabetic subjects have attenuated exercise-stimulated AMPK activity, and the authors (65) suggest that obese type 2 diabetic subjects need to exercise at higher intensity to stimulate AMPK phosphorylation. Since type 2 diabetes mellitus is strongly correlated with obesity and with a sedentary lifestyle, a weak activation of AMPK in the periphery, due to over-eating and/or lack of exercise, may be a contributory factor in its onset.

There are species differences in the adaptive response of skeletal muscle to food deprivation. The data described above clearly show, in the rat, an active adaptive response to food deprivation through a regulation of skeletal muscle AMPK activity by fatty acid-mediated signaling. Surprisingly, no such relation between food deprivation and skeletal muscle AMPK phosphorylation has been found in mice (66) . However, since in that study AMPK phosphorylation was measured only at a single time point (24 h after food deprivation), the transient stimulation of AMPK phosphorylation may have been missed. In favor of this assumption, a functional role for AMPK in triggering lipid metabolism during fasting has been demonstrated in mice, since AMPK {gamma}3 subunit knockout mice starved for 16 h failed to up-regulate genes controlling glucose and lipid metabolism (lipoprotein lipase, fatty acid translocase, carnitine palmitoyl transferase 1b, and citrate synthase; ref. 56 ). Further, DNA microarray studies performed on skeletal muscle obtained from 24 and 48 h fasted mice (7) and 48 h fasted rats (6) have clearly shown that structural/metabolic adaptations to fasting at the transcriptional level are more pronounced in the rat than in the mouse. An increase in myosin heavy chain I gene expression, leading to type I fiber formation (a process that is PGC1-{alpha}/PPAR{delta} mediated) (33 , 34) is not observed in fasted-mouse skeletal muscle, and the increases seen in rat skeletal muscle in the very long, long, and medium Acyl CoA dehydrogenases (VLCAD, LCAD, and MCAD) (6) , all of them PPAR{delta} targets (31) , are not seen in gastrocnemius muscle from mice deprived of food for 24 or 48 h (7) . At these time points, transcriptional changes in the fasted mouse muscle focus on suppressing glucose oxidation/protein degradation as well as reduced energy uncoupling and include atrophy-specific changes in gene expression (7) . In the fasted rat, the mRNA levels of several genes involved in lipid metabolism that were initially up-regulated are already down-regulated after 48 h (6) , and it may be that this is also true for mouse mRNA levels at the same time-points. Indeed, the work cited above by Long et al. (56) showed that in mice starved for 16 h, genes controlling lipid metabolism are up-regulated. A more complete time-course study of food deprivation in the mouse has proven to be more informative. In starved mice, the mRNA for a member of the FOXO family, forkhead homologue in rhabdomyosarcoma (FKHR), was up-regulated rapidly and transiently (starting within 6 h, peaking at 12 h, and decreasing at 24 h), before the up-regulation of FKHR protein levels (and relatively low phosphorylation at position Ser-256), with a consequent up-regulation of its target gene pyruvate dehydrogenase kinase 4 (PDK4) (67) . This kinase plays an important role in the switching from glucose to fat usage during starvation since it phosphorylates the E1 component of the pyruvate dehydrogenase (PDH) complex, thereby down-regulating carbohydrate (CH) oxidation (68) . Similarly, single (or dual) time point studies have not yielded clear conclusions on the adaptive changes in human skeletal muscle with respect to the fasting-induced increase in lipid metabolism. Upon 24 or 48 h of fasting in 10 healthy human male subjects, despite the elevated plasma FFA levels, no altered expressions were seen for transcription factors, MHC genes, or genes involved in fat metabolism (45) . Interestingly, in analogy to what has been reported in the mouse (67) , PDK4 mRNA and protein levels were elevated at both 24 and 48 h; however, at these time points, neither the FOXO1 mRNA/protein levels nor FOXO1 phosphorylation at Ser-256 showed any change (45) . Nevertheless, it remains a possibility that regulation of the level of the mRNA/protein of FOXO1 (a PGC-1{alpha} target) (19) are transient events that precede up-regulation of PDK4 mRNA and protein and that they may therefore have been missed by measuring only at 24 and 48 h of starvation in humans. At the 48 h time point, PPAR{delta} mRNA levels in the human vastus lateralis muscle were decreased (45) , which is in line with the decrease in the levels of PPAR {delta} mRNA (6) and nuclear protein (13) at this time point in rat gastrocnemius muscle. In the rat, this decline was preceded by a rapid rise in the mRNA and protein levels of this transcription factor at 6 and 12 h of food deprivation (13) , but these time points were not investigated in the human study. Alternate day fasting in humans (for 3 wk) did not result in evidence of alterations in skeletal muscle gene expression in human vastus lateralis muscle (69) .

To clarify the apparent discrepancies among species, additional research, including more comprehensive time-course studies, needs to be carried out. With regard to the timing of the changes in the expression/phosphorylation of some important factors involved in the primary adaptative response of skeletal muscle to fasting, a comparison among mice, rats, and humans is given in Fig. 3 .


Figure 3
View larger version (28K):
[in this window]
[in a new window]

  		Figure 3. Timing of expression/phosphorylation of several important factors involved in primary adaptative response of skeletal muscle to food deprivation: comparison among rats, mice, and humans. Reference numbers from which the data were obtained are indicated above histograms. N.D. = not determined in any of investigated species.

NO has a role as a signaling molecule during food deprivation. NO is a free radical produced from L-arginine by various isoforms of NO synthase (NOS) in virtually all cell-types. At pathological levels NO acts as an oxidant causing the damaging effects typical for free radicals. At physiological levels, however, NO acts as a signaling molecule to stimulate glucose uptake as well as the oxidation of glucose and fatty acids in skeletal muscle, heart, liver, and adipose tissue (see, for review, ref. 40 ). Recently, overexpression of endothelial NOS (eNOS) has been shown to affect the expression levels of PGC-1{alpha}. Thus, NO produced by eNOS activates guanylate cyclase to increase the amount of cyclic GMP (cGMP) present, transmitting a signal to the nucleus that causes the induction of PGC-1{alpha}-gene transcription (70) . The same group (71) showed that life span-increasing caloric restriction in mice (for either 3 or 12 months) induced eNOS expression in various tissues, including the heart. The induced eNOS correlated with the increased PGC-1{alpha} expression and with mitochondrial biogenesis, increased oxygen consumption, and ATP production, and also with an enhanced expression of the yeast silent information regulator 2 (Sir2) homologue SIRT1 (71) . Sir2-related enzymes, generally termed sirtuins, are a recently discovered class of NAD(+)-dependent protein deacetylases that regulate gene expression in a variety of organisms by deacetylation of modified lysine residues on histones, transcription factors, and other proteins, and are identified as important regulators of organism life span (for review, see ref. 72 ). Very recently, it has been shown that SIRT1 deacetylates PGC-1{alpha} under low glucose conditions in skeletal muscle cells (with concomitant elevation of NAD+ levels), inducing a metabolic gene transcription program of mitochondrial fatty acid oxidation (73) , although the authors did not address the involvement of NO-mediated signaling in this process. Interestingly, a natural polyphenolic compound found in the skin of grapes known to increase life span, resveratrol, has been shown to improve mitochondrial function by inducing SIRT1/PGC-1{alpha} signaling as well as a shift toward more oxidative fiber types (without a complete shift toward type I fibers) (74) . Again, the role of NO in the effect of resveratrol has not been investigated. Intriguingly, mitochondria can induce their own synthesis: a mitochondrial SIRT isoform, SIRT3, has been shown to promote mitochondrial biogenesis through increased PGC-1{alpha} expression in the brown adipose tissue of calorie-restricted mice (through a currently unknown mechanism) (75) . Whether (and when) NO operates through SIRT signaling to induce lipid metabolism in skeletal muscle during fasting is still an open question. Up-regulation of SIRT 1 expression, without changes in the expression of genes involved in fatty acid transport/oxidation or mitochondrial biogenesis, has been demonstrated in long-term fasted human skeletal muscle (69) . In a recent study, NO metabolites were measured in the plasma of rats submitted to food deprivation for 3 days, the results suggesting that fasting leads to an increased plasma NO level, probably by activation of NOS (76) . However, the kinetics of this process during fasting remains unknown.

AMPK- and NO-mediated signaling are two intertwining pathways. AMPK plays a stimulatory role in NO production. Moreover, AMPK phosphorylates the predominant NOS isoform, nNOSµ, in human skeletal muscle during exercise (77) , indicating a role for NO in AMPK-mediated signaling during exercise-induced energy deprivation. Indeed, blocking NOS activity in human isolated skeletal muscle diminishes AICAR-stimulated glucose uptake (77) , although experiments on isolated rat epitrochlearis muscle subjected to contractions did not show reduced AICAR-induced glucose uptake upon NOS inhibition [induced by administration of nitro-L-arginine methyl ester (L-NAME)], suggesting pathways other than AMPK-NOS may be involved in contraction-induced glucose uptake (78) . Not only can AMPK regulate NO synthesis, but NO can modulate AMPK activity. It has recently been shown that a product of the spontaneous reaction of NOwith superoxide anion (O2–), namely peroxynitrite (ONOO–), activates AMPK through a c-Src-mediated and phosphatidylinositol 3-kinase (PI3K)-dependent pathway in various tissues, including mouse heart, without a change in the AMP/ATP ratio (79) . Importantly, it has been shown in vivo that NO increases AMPK-induced glucose uptake but not long chain fatty acid (LCFA) uptake in rat skeletal muscle, with the LCFA uptake increasing in type I fibers (80) . This shows that the mechanisms by which AMPK stimulates glucose and LCFA uptake are distinct from each other and moreover that these data would imply that NO is a less important factor in the regulation of energy homeostasis during fasting. Taking all these data into account, it is clear that the role performed by NO in the process elicited by AMPK during fasting needs further investigation.


   SKELETAL MUSCLE MITOCHONDRIA AND FOOD DEPRIVATION
TOP
 ABSTRACT
 INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 
The increased mitochondrial biogenesis and increased FFA oxidation seen during food deprivation are induced by AMPK. Since type I fibers are slow twitch, with a high capacity for the synthesis of ATP and many mitochondria that actively oxidize fatty acids, it is perhaps not surprising that the above-mentioned AMPK-induced factors involved in the switching of muscle fibers toward an oxidative phenotype also stimulate mitochondrial biogenesis. It is well known that PGC-1{alpha} is a master regulator of mitochondrial biogenesis (19) , and moreover PPAR{delta} has been shown to induce the expression of genes involved in this process (31 ; for a detailed review on mitochondrial biogenesis induced by these factors, see ref. 32 ). In food-deprived skeletal muscle, mitochondrial fatty acid-oxidizing activity increases (13) . Acetyl-CoA, a common intermediate of glucose, fatty acid, and amino acid oxidation, either enters the Krebs cycle for complete oxidation or is utilized for lipogenesis in skeletal and cardiac muscles, depending on the intracellular energy status and activity level of acetyl-CoA carboxylase (ACC) (81) . In skeletal and cardiac muscles, as in the liver, malonyl CoA, a product of the ACC reaction, is an allosteric inhibitor of carnitine palmitoyl transferase 1 (CPT 1), the enzyme that transfers the long-chain fatty acyl-CoA into mitochondria for ß oxidation (82) . Reduced levels of malonyl-CoA favor LCFA oxidation (82 , 83) . This process has long been known to occur during fasting, but only recently has it been shown that during food deprivation AMPK phosphorylates and inactivates ACC at position Ser-79, reducing malonyl CoA levels and increasing mitochondrial LCFA oxidation in skeletal muscle (13 ; see Fig. 1 in the present review). Another level of regulation of fatty acid utilization by AMPK, other than by the AMPK-ACC-CPT pathway, involves the uptake of LCFA through the sarcolemma. Indeed, during fasting the greater supply of LCFA for muscle-cell oxidation seems to be mediated by membrane enrichment with fatty acid translocase/CD36m (FAT/CD36). FAT/CD36 mRNA is up-regulated during 24 h starvation, while refeeding promotes down-regulation of its gene (46) . Moreover, acute regulation of LCFA uptake by muscle cells involves cellular redistribution from an intracellular compartment to the surface of the sarcolemma (84 , 85) , a process that is regulated by AMPK and induced by AICAR in cardiomyocytes (85) . The combined induction of FAT/CD36 mobilization and CPT-1 activity though AMPK activation is a metabolically efficient process allowing the extra incoming fatty acid to be preferentially channeled toward mitochondrial beta-oxidation. The FAT/CD36 protein is also located within the inner mitochondrial membrane (86) and is postulated to receive the lipid-binding protein (LPB) bound LCFAs from the cytosol to hand them off to the long chain acyl CoA synthetase (ACS) that activates them (86) , being subsequently transported into the matrix by the CPT system that is activated by AMPK.

There is increased UCP3 expression during fasting that relates to the role of AMPK. In conditions of altered metabolism, such as fasting, the UCP predominantly expressed in skeletal muscle, UCP3, may play a role in mitochondrial lipid handling. Indeed, the effects of physiological interventions consistently support a role for UCP3 either in promoting fatty acid oxidation (87 , 88) or in the translocation of peroxidised fatty acids (89) . During fasting, the expression of uncoupling protein 3 is increased through several pathways, which may possibly interact. These include: PPAR {delta} signaling (31) , the AMPK pathway (10 11 12) , and MyoD, an essential factor for skeletal muscle UCP3 transcription (90) , operating in functional association with PPAR signaling (91) and activated by muscle LIM (Lin-11, Isl-1, Mec-3, representing the genes in which the motif was originally discovered) protein (92) . This protein contains the LIM domain, an evolutionary conserved double-zinc finger motif found in a variety of proteins exhibiting diverse biological roles. LIM domains have been observed to act as modular protein-binding interfaces mediating protein-protein interactions in the cytoplasm and the nucleus. Several LIM domain-containing proteins associated with the actin cytoskeleton have been identified, playing a role in signal transduction and organization of the actin filaments during various cellular processes (see, for review, ref. 93 ). The expression of muscle LIM protein is increased dramatically during fasting (6) . Numerous reports have proposed an interplay between UCP3 and AMPK, with AMPK activity being a trigger for UCP3 expression (10 11 12) or vice versa (94) . Recently, we presented evidence that in the gastrocnemius muscle of food-deprived rats, increased AMPK activity is unlikely to be the consequence of an increased expression of UCP3, since AMPK phosphorylation preceded the increase in UCP3 expression (13) . Indeed, our results imply that the increases in mRNA levels and mitochondrial protein levels of UCP3 are events downstream of AMPK phosphorylation. These data led us to conclude that UCP3 is not responsible for the increased phosphorylation of AMPK but rather that fluctuating levels of fatty acids are most likely involved in AMPK activation/deactivation in skeletal muscle. A visual overview of AMPK signaling related to increased mitochondrial biogenesis through SIRT/PGC-1{alpha}, increased UCP3 expression, decreased carbohydrate oxidation, and increased fatty acid transport/oxidation in food-deprived skeletal muscle is given in Fig. 4 .


Figure 4
View larger version (42K):
[in this window]
[in a new window]

  		Figure 4. Consequences of AMPK phosphorylation in food-deprived skeletal muscle cell. Increased mitochondrial biogenesis through SIRT/PGC-1{alpha}, increased UCP3 expression, decreased carbohydrate oxidation, and increased fatty acid transport/oxidation. White arrow pointing upward depicts a stimulatory effect; white arrow pointing downward depicts an inhibitory effect. All other arrows depict stimulatory effects. Question mark (?) depicts a currently unknown mechanism. Triangles represent free fatty acid molecules. For all abbreviations and for further explanations, refer to text.


   CONCLUSIONS AND FUTURE PROSPECTS
TOP
 ABSTRACT
 INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
REFERENCES
 
Recent observations have revealed that food deprivation, like exercise (for a recent review, see ref. 95 ), represents a physiological situation in which the fuel-providing and regulatory role of AMPK in skeletal muscle is prominent (see Figs. 1 and 4 , and for a schematic overview, Fig. 5 ). Food deprivation and exercise share the same kinetics of activation of the signaling pathways triggering structural/metabolic adaptations toward lipid metabolism in skeletal muscle. AMPK activation leads to increased FFA levels and an increased mitochondrial uptake of FFA over glucose as the fuel substrate. This process seems to be NO independent but AMP dependent. In turn, the increased intracellular FFA levels, through an inhibition of AMPK phosphorylation, guarantee the sparing of glucose for use by the brain (for a temporal scheme of the events, see Fig. 2 ). Randle et al. (96) first used the term "glucose-fatty acid cycle" to describe the reciprocal inhibition of glucose metabolism by high-level fatty acid utilization in the heart. Besides inactivation of the pyruvate dehydrogenase complex during fasting, inhibition of glucose uptake is also achieved by inhibiting AMPK phosphorylation. An intriguing, but still open, question is how do fatty acids (or, eventually, their metabolic products) bring about inhibition of AMPK phosphorylation in this condition? It is well known that increases in fatty acids mediate inhibition of insulin signaling in skeletal muscle through phosphorylation of nPKC via increased levels of diacylglycerol (97) , which in turn stimulates serine phosphorylation and inhibits tyrosine phosphorylation and activation of insulin receptor substrates (IRS) (97) . It is conceivable that fatty acids act through a yet unidentified protein kinase to counteract the effect of LKB1 or other upstream activators of AMPK.


Figure 5
View larger version (19K):
[in this window]
[in a new window]

  		Figure 5. General overview of the structural metabolic events taking place in skeletal muscle after food deprivation.

There are many indications that mice and rats differ in their responses to food deprivation. It seems increasingly clear that rats have developed a much more versatile strategy to enable them to survive the stress of food deprivation, essentially through a remodeling of their muscle mass toward fatty acid metabolism. Further, given the rapid kinetics of fasting-induced AMPK-mediated signaling, studies on mice and humans that involve only single time point measurements may not be conclusive due to the lack of time course data. However, such data are especially difficult to obtain in human subjects.


   ACKNOWLEDGMENTS
 
This study was supported in part by MIUR-COFIN 2003 protocol no. 2003052503–002.

Received for publication March 12, 2007. Accepted for publication May 17, 2007.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 FACTORS INVOLVED IN STRUCTURAL...
 MAINTENANCE OF SKELETAL MUSCLE...
 SKELETAL MUSCLE MITOCHONDRIA AND...
 CONCLUSIONS AND FUTURE PROSPECTS
 REFERENCES
 

  1. Fluck, M., Hoppeler, H. (2003) Molecular basis of skeletal muscle plasticity—from gene to form and function. Rev. Physiol. Biochem. Pharmacol. 146,159-216[Medline]
  2. Pette, D., Staron, R. S. (1990) Cellular and molecular diversities of mammalian skeletal muscle fibers. Rev. Physiol. Biochem. Pharmacol. 116,1-76[Medline]
  3. Pette, D. (2002) The adaptive potential of skeletal muscle fibers. Can. J. Appl. Physiol. 27,423-448[Medline]
  4. Kyrolainen, H., Kivela, R., Koskinen, S., McBride, J., Andersen, J. L., Takala, T., Sipila, S., Komi, P. V. (2003) Interrelationships between muscle structure, muscle strength, and running economy. Med. Sci. Sports. Exerc. 35,45-49
  5. Pilegaard, H., Saltin, B., Darrell Neufer, P. (2003) Effect of short term fasting and refeeding on transcriptional regulation of metabolic genes in human skeletal muscle. Diabetes 52,657-662[Abstract/Free Full Text]
  6. De Lange, P., Ragni, M., Silvestri, E., Moreno, M., Schiavo, L., Lombardi, A., Farina, P., Feola, A., Goglia, F., Lanni, A. (2004) Combined cDNA array/RT PCR analysis of gene expression profile in rat gastrocnemius muscle: relation to its adaptive function in energy metabolism during fasting. FASEB J. 18,350-352[Abstract/Free Full Text]
  7. Jagoe, R. T., Lecker, S. R., Gomes, M., Goldberg, A. L. (2002) Patterns of gene expression in atrophying skeletal muscle: response to food deprivation. FASEB J. 16,1697-1712[Abstract/Free Full Text]
  8. Hardie, D. G., Hawley, S. A., Scott, J. W. (2006) AMP-activated protein kinase—development of the energy sensor concept. J. Physiol. 574,7-15[Abstract/Free Full Text]
  9. Hardie, D. G., Carling, D., Carlson, M. (1998) The AMP activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryote cell?. Annu. Rev. Biochem. 67,821.855
  10. Zhou, M., Lin, B.-Z., Coughlin, S., Vallega, G., Philch, P. F. (2000) UCP-3 expression in skeletal muscle: effects of exercise, hypoxia, and AMP-activated protein kinase. Am. J. Physiol. Endocrinol. Metab. 279,622-629
  11. Suwa, M., Nakano, H., Kugami, S. (2003) Effects of chronic AICAR treatment on fiber composition, enzyme activity, UCP3, and PGC-1 in rat muscles. J. Appl. Physiol. 95,950-958
  12. Putman, C. T., Kricsi, M., Pearcy, J., MacLean, I. M., Bamford, J. A., Murdoch, G. K., Dixon, W. T., Pette, D. (2003) AMPK activation increases uncoupling protein-3 expression and mitochondrial enzyme activities in rat muscle without fibre type transitions. J. Physiol. 551,169-178[Abstract/Free Full Text]
  13. De Lange, P., Farina, P., Moreno, M., Ragni, M., Lombardi, A., Silvestri, E., Burrone, L., Lanni, A., Goglia, F. (2006) Sequential changes in the signal transduction responses of skeletal muscle following food deprivation. FASEB J. 20,2579-2581[Abstract/Free Full Text]
  14. Jaschinski, F., Schuler, M., Peuker, H., Pette, D. (1998) Changes in myosin heavy chain mRNA and protein isoforms of rat muscle during forced contractile activity. Am. J. Physiol. Cell Physiol. 274,C365-C370[Abstract/Free Full Text]
  15. Spiegelman, B. M., Heinrich, R. (2004) Biological control through regulated transcriptional coactivators. Cell 119,157-167[CrossRef][Medline]
  16. Knutti, D., Kaul, A., Tralli, A. (2000) A tissue-specific coactivator of steroid receptors, identified in a functional genetic screen. Mol. Cell. Biol. 20,2411-2422[Abstract/Free Full Text]
  17. Lin, J., Puigserver, P., Donovan, J., Tarr, P., Spiegelman, B. M. (2002) Peroxisome proliferator-activated receptor gamma coactivator 1beta (PGC-1 beta), a novel PGC-1 related transcription coactivator associated with host cell factor. J. Biol. Chem. 277,1645-1648[Abstract/Free Full Text]
  18. Puigserver, P., Wu, Z., Park, C. W., Graves, R., Wright, M., Spiegellman, B. M. (1998) A cold-inducible coactivator of nuclear receptors is linked to adaptive thermogenesis. Cell 92,829-839[CrossRef][Medline]
  19. Lin, J., Handschin, C., Spiegelman, B. M. (2005) Metabolic control through the PGC-1 family of transcription coactivators. Cell Metab. 1,361-370[CrossRef][Medline]
  20. Wu, Z., Puigserver, P., Andersson, U., Zhang, C., Adelmant, G., Mootha, V., Troy, A., Cinti, S., Lowell, B., Scarpulla, R. C., et al (1999) Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell 98,115-124[CrossRef][Medline]
  21. Vega, R. B., Huss, J. M., Kelly, D. P. (2000) The coactivator PGC-1 cooperates with peroxisome proliferators-activated receptor alpha in transcriptional control of nuclear genes encoding mitochondrial fatty acid oxidation enzymes. Mol. Cell. Biol. 20,1868-1876[Abstract/Free Full Text]
  22. Wang, Y. X., Lee, C. H., Tiep, S., Yu, R. T., Ham, J., Kang, H., Evans, R. M. (2003) Peroxisome-proliferator-activated receptor {delta} activates fat metabolism to prevent obesity. Cell 113,159-170[CrossRef][Medline]
  23. Huss, J. M., Kopp, R. P., Kelly, D. P. (2002) Peroxisome proliferators-activated receptor coactivator-1alpha (PGC-1alpha) coactivates the cardiac-enriched nuclear receptors estrogen-related receptor-alpha and gamma. Identification of novel leucine-rich interaction motif within PGC-1alpha. J. Biol. Chem. 277,40265-40274[Abstract/Free Full Text]
  24. Huss, J. M., Torra, I. P., Staels, B., Giguere, V., Kelly, D. P. (2004) Estrogen-related receptor alpha directs peroxisome proliferator-activated receptor alpha signaling in the transcriptional control of energy metabolism in cardiac and skeletal muscle. Mol. Cell. Biol. 24,9079-9091[Abstract/Free Full Text]
  25. Kamei, Y., Ohizumi, H., Fujitani, Y., Remoto, T., Tanaka, T., Takahashi, N., Kawada, T., Miyoshi, M., Ezaki, O., Kakizuka, A. (2003) PPAR gamma coactivator 1 beta/ERR ligand 1 is an ERR protein ligand, whose expression induces a high-energy expenditure and antagonizes obesity. Proc. Natl. Acad. Sci. U. S. A. 100,12378-12383[Abstract/Free Full Text]
  26. Mootha, V. K., Handschin, C., Arlow, D., Xie, X., St. Pierre, J., Sihag, S., Yang, W., Altshuler, D., Puigserver, P., Pattersom, N., et al (2004) Err(alpha) and Gabpa/b specify PGC-1 (alpha)-dependent oxidative phosphorylation gene expression that is altered in diabetic muscle. Proc. Natl. Acad. Sci. U. S. A. 101,6570-6575[Abstract/Free Full Text]
  27. Schreiber, S.N., Emter, R., Hock, M. B., Knutti, D., Cardenas, J., Podvinec, M., Oakeley, E. J., Kralli, A. (2004) The estrogen-related receptor {alpha} (ERR{alpha}) functions in PPAR{gamma} coactivator 1{alpha} (PGC-1{alpha})-induced miochondrial biogenesis. Proc. Natl. Acad. Sci. U. S. A. 101,6472-6477[Abstract/Free Full Text]
  28. Zhang, Y., Ma, K., Song, S., Elam, M. B., Cook, G. A., Park, E. A. (2004) Peroxisomal proliferators-activated receptor gamma coactivator-1 alpha (PGC-1alpha) enhances the thyroid hormone induction of carnitine palmitoyl transferase I (CPT-I alpha). J. Biol. Chem. 279,53963-53971[Abstract/Free Full Text]
  29. Kelly, D. P., Scarpulla, R. C. (2004) Transcriptional regulatory circuits controlling mitochondrial biogenesis and function. Genes Dev. 18,357-368[Free Full Text]
  30. Puigserver, P., Spiegelman, B. M. (2003) Peroxisome proliferators-activated receptor-gamma coactivator 1 alpha (PGC-1alpha): transcriptional coactivator and metabolic regulator. Endocr. Rev. 24,78-90[Abstract/Free Full Text]
  31. Barish, G. D., Narkar, V. H., Evans, R. M. (2006) PPAR{delta}: a dagger in the heart of the metabolic syndrome. J. Clin. Invest. 116,590-597[CrossRef][Medline]
  32. Reznick, R. M., Shulman, G. I. (2006) The role of AMP-activated protein kinase in mitochondrial biogenesis. J. Physiol. 574,33-39[Abstract/Free Full Text]
  33. Lin, J., Wu, H., Tarr, P. T., Zhang, C.-Y., Wu, Z., Boss, O., Michael, L. F., Puigserver, P., Isotani, E., Olson, E. N., et al (2002) Transcriptional co-activator PGC-1{alpha} drives the formation of slow-twitch muscle fibers. Nature 419,797-801[CrossRef][Medline]
  34. Wang, Y.X., Zhang, C.-L., Yu, R. T., Cho, H. K., Nelson, M. C., Bayugu-Ocampo, C. R., Ham, J., Kang, H., Evans, R. M. (2004) Regulation of muscle fiber-type and running endurance by PPAR{delta}. PloS. Biol. 2,e294[CrossRef][Medline]
  35. Leone, T. C., Lehman, J. J., Finck, B. N., Schaeffer, P. J., Wende, A. R., Boudina, S., Courtois, M., Wozniak, D. F., Sambandam, N., Bernal-Mizrachi, C., et al (2005) PGC-1alpha deficiency causes multi-system energy metabolic derangements: muscle dysfunction, abnormal weight control and hepatic steatosis. PloS Biol. 3,e101[CrossRef][Medline]
  36. Vescovo, G., Ravara, B., Gobbo, V., Angelini, A., Dalla Libera, L. (2005) Skeletal muscle fibres synthesis in heart failure: role of PGC-1alpha, calcineurin and GH. Int. J. Cardiol. 104,298-306[CrossRef][Medline]
  37. Irrcher, I., Adhihetty, P. J., Sheehan, T., Joseph, A. M., Hood, D. A. (2003) PPAR gamma coactivator-1 alpha expression during thyroid hormone- and contractile activity- induced mitochondrial adaptations. Am. J. Physiol. Cell. Physiol. 284,C1699-C1677
  38. Terada, S., Goto, M., Kato, M., Kawanaka, K., Shimokawa, T., Tabata, I. (2002) Effects of low-intensity prolonged exercise on PGC-1 mRNA expression in rat epitrochlearis muscle. Biochem. Biophys. Res. Commun. 296,350-354[CrossRef][Medline]
  39. Zong, H., Ren., J. M., Young, L. H., Pypaert, M., Mu, J., Birnbaum, M. J., Shulman, G. I. (2002) AMP kinase is required for mitochondrial biogenesis in skeletal muscle in response to chronic energy deprivation. Proc. Natl. Acad. Sci. U. S. A. 25,15983-15987
  40. Jobgen, W. S., Fried, S. K., Fu, W. J., Meininger, C. J., Wu, G. (2006) Regulatory role for the arginine-nitric oxide pathway in metabolism of energy substrates. J. Nutr. Biochem. 17,571-588[CrossRef][Medline]
  41. Escher, P., Braissant, O., Basu-Modak, S., Michalik, L., Wahli, W., Desvergne, B. (2001) Rat PPARs: Quantitative analysis in adult rat tissues and regulation in fasting and refeeding. Endocrinology 142,4195-4202[Abstract/Free Full Text]
  42. Luquet, S., Lopez-Soriano, J., Holst, D., Fredenrich, A., Malki, J., Rassoulzadegan, M., Grimaldi, P. A. (2003) Peroxisome proliferator-activated receptor {delta} controls muscle development and oxidative capability. FASEB J. 17,2299-2301[Abstract/Free Full Text]
  43. Brunmair, B., Staniek, K., Dorig, J., Szocs, Z., Stadlbauer, K., Marian, V., Gras, F., Anderwald, C., Nohl, H., Waldhsausl, W., et al (2006) Activation of PPAR-delta in isolated rat skeletal muscle switches fuel preference from glucose to fatty acids. Diabetologia 49,2713-2722[CrossRef][Medline]
  44. Tanaka, T., Yamamoto, J., Iwasaki, S., Asaba, H., Hamura, H., Ikeda, Y., Watanabe, M., Magoori, K., Ioka, R. X., Tachibana, K., et al (2003) Activation of peroxisome proliferators-activated receptor {delta} induces fatty acid ß oxidation in skeletal muscle and attenuates metabolic syndrome. Proc. Natl. Acad. Sci. U. S. A. 100,15924-15929[Abstract/Free Full Text]
  45. Tsintzas, K., Jewell, K., Kamran, M., Laithwaite, D., Boonsong, T., Littlewood, J., MacDonald, I., Bennet, A. (2006) Differential regulation of metabolic genes in skeletal muscle during starvation and refeeding in humans. J. Physiol. 575,291-303[Abstract/Free Full Text]
  46. Holst, D., Luquet, S., Nogueira, V., Kristiansen, K., Leverve, X., Grimaldi, P. A. (2003) Nutritional regulation and role of peroxisome proliferators-activated receptor delta in fatty acid catabolism in skeletal muscle. Biochim. Biophys. Acta 1633,43-50[Medline]
  47. Stavinoha, M. A., RaySpellicy, J. W., Essop, M. F., Graveleau, C., Abel, E. D., Hart-Sailors, M. L., Mersmann, H. J., Bray, M. S., Young, M. E. (2004) Evidence for mitochondrial thioesterase 1 as a peroxisome proliferator-activated receptor alpha-regulated gene in cardiac and skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 287,E888-E895[Abstract/Free Full Text]
  48. Watt, M. J., Southgate, R. J., Holmes, A. G., Febbraio, M. A. (2004) Suppression of plasma free fatty acids upregulates peroxisome proliferators-activated (PPAR) alpha and delta and PPAR coactivator 1 alpha in human skeletal muscle, but not lipid regulatory genes. J. Mol. Endocrinol. 33,533-544[Abstract/Free Full Text]
  49. Mahoney, D. J., Parise, G., Melov., S., Safidar, A., Tarnopolky, M. A. (2005) Analysis of global mRNA expression in human skeletal muscle during recovery from endurance exercise. FASEB J. 19,1498-1500[Abstract/Free Full Text]
  50. Hawley, S.A., Boudeau, J., Reid, J. L., Mustard, K. J., Udd, L., Makela, T. P., Alessi, D. R., Hardie, D. G. (2003) Complexes between the LKB1 tumor suppressor, STRAD {alpha}/ß and MO25 {alpha}/ß are upstream kinases in the AMP-activated protein kinase cascade. J. Biol. 2,28[CrossRef][Medline]
  51. Sakamoto, K., Zarrinpashneh, E., Budas, G. R., Pouleur, A. C., Dutta, A., Prescott, A. R., Ashworth, A., Jovanovic, A., Alessi, D. R., Bertrand, L. (2006) Deficiency of LKB1 in heart prevents ischemia-mediated activation of AMPK{alpha}2 but not AMPK{alpha}1. Am. J. Physiol. Endocrinol. Metab. 290,E780-E788[Abstract/Free Full Text]
  52. Lizcano, J. M., Goransson, O., Toth, R., Deak, M., Morrice, N. A., Boudeau, J., Hawley, S. A., Udd, L., Makela, T. P., Hardie, D. G., et al (2004) LKB is a master kinase that activates 13 protein kinases of the AMPK subfamily, including the MARK/PAR-1 kinases. EMBO J. 23,833-843[CrossRef][Medline]
  53. Sakamoto, K., Goransson, O., Hardie, D. G., Alessi, D. R. (2004) Activity of LKB1 and AMPK-related kinases in skeletal muscle: effects of contraction, phenformin, and AICAR. Am. J. Physiol. Endocrinol. Metab. 287,E310-E317[Abstract/Free Full Text]
  54. Winder, W. W., Hardie, D. G., Mustard, K.J., Greenwood, L. J., Paxton, B. E., Park, S. H., Rubink, D. S., Taylor, E. B. (2003) Long-term regulation of AMP-activated protein kinase and acetyl-CoA carboxylase in skeletal muscle. Biochem. Soc. Trans. 31,182-185[Medline]
  55. Park, H., Kaushik, V. K., Constant, S., Prentki, M., Przybytkowski, E., Ruderman, N. B., Saha, A. K. (2002) Coordinate regulation of malonyl-CoA decarboxylase, sn-glycerol-3-phosphate acyltransferase, and acetyl-CoA carboxylase by AMP-activated protein kinase in rat tissues in response to exercise. J. Biol. Chem. 277,32571-32577[Abstract/Free Full Text]
  56. Long, Y. C., Barnes, B. R., Mahlapuu, M., Steiler, T. L., Martinsson, S., Leng, Y., Wallberg-Hendriksson, H., Andersson, L., Zierath, J. R. (2005) Role of AMP-activated protein kinase in the coordinated expression of genes controlling glucose and lipid metabolism in mouse white skeletal muscle. Diabetologia 48,2354-2365[CrossRef][Medline]
  57. Jorgensen, S. B., Wojtaszewski, J. F., Viollet, B., Andreelli, F., Birk, J. B., Hellsten, Y., Schjerling, P., Vaulont, S., Neufer, P. D., Richter, E. A., et al (2005) Effects of {alpha}-AMPK knockout on exercise-induced gene activation in mouse skeletal muscle. FASEB J. 19,1146-1148[Abstract/Free Full Text]
  58. Itani, S. I., Saha, A. K., Kurowski, T. G., Coffin, H. R., Tornheim, K., Ruderman, N. B. (2003) Glucose autoregulates its uptake in skeletal muscle. Diabetes 53,1635-1640
  59. Jing, M., Ismail-Beigi, F. (2007) Critical role of 5'-AMP-activated protein kinase in the stimulation of glucose transport in response to inhibition of oxidative phosphorylation. Am. J. Physiol. Cell. Physiol. 292,C477-C487[Abstract/Free Full Text]
  60. Lin, J., Wu, P. H., Tarr, P. T., Lindenberg, K. S., St-Pierre, J., Zhang, W., Mootha, V. K., Jager, S., Vianna, C. R., Reznick, R. M., et al (2004) Defects in adaptative energy metabolism with CNS-linked hyperactivity in PGC-1alpha null mice. Cell 119,121-135[CrossRef][Medline]
  61. Sriwijitkamol, A., Ivy, J. L., Christ-Roberts, C., Defronzo, R. A., Mandarino, L. J., Musi, N. (2006) LKB1-AMPK signaling in muscle from obese insulin-resistant Zucker rats and effects of training. Am. J. Physiol. Endocrinol. Metab. 290,E925-E932[Abstract/Free Full Text]
  62. Liu, Y., Wan, Q., Guan, Q., Gao, L., Zhao, J. (2006) High-fat diet feeding impairs both the expression and activity of AMPK{alpha} in rats’ skeletal muscle. Biochem. Biophys. Res. Commun. 339,701-707[CrossRef][Medline]
  63. Hojlund, K., Mustard, K. J., Staehr, P., Hardie, D. G., Beck-Nielsen, H., Richter, E. A., Wojtaszweski, J. F. (2004) AMPK activity and subunit protein expression are similar in muscle of obese subjects with and without type 2 diabetes. Am. J. Physiol. Endocrinol. Metab. 286,E239-E244[Abstract/Free Full Text]
  64. Steinberg, G. R., Smith, A. C., Van Denderen, B. J., Chen, Z., Murthy, S., Campbell, D. J., Heigenhauser, G. J., Dyck, D. J., Kemp, B. E. (2004) AMP-activated protein kinase is not down-regulated in human skeletal muscle of obese females. J. Clin. Endocrinol. Metab. 89,4575-4580[Abstract/Free Full Text]
  65. Sriwijitkamol, A., Coletta, D. K., Wajcberg, E., Balbontin, G. B., Reyna, S. M., Barrientes, J., Eagan, P. A., Jenkinson, C. P., Cersosimo, E., Defronzo, R. A., et al (2007) Effect of acute exercise on AMPK signaling in skeletal muscle of subjects with type 2 diabetes: A time-course and dose-response study. Diabetes 56,836-848[Abstract/Free Full Text]
  66. Gonzalez, A. A., Kumar, R., Mulligan, J. D., Davis, A. J., Weindruch, R., Saupe, K. (2004) Metabolic adaptations to fasting and chronic caloric restriction in heart, muscle and liver do not include changes in AMPK activity. Am. J. Physiol. Endocrinol. Metab. 287,R1270-R1275
  67. Furuyama, T., Kitamaya, K., Yamashita, H., Mori, N. (2003) Forkhead transcription factor FOXO1 (FKHR)-dependent induction of PDK4 gene expression in skeletal muscle during energy deprivation. Biochem. J. 375,365-371[CrossRef][Medline]
  68. Sugden, M. C., Howard, R. M., Munday, M. R., Holness, M. J. (1993) Mechanisms involved in the coordinated regulation of strategic enzymes of glucose metabolism. Ad. Enzyme Regul. 33,71-95[CrossRef]
  69. Heilbronn, L. K., Civitarese, A. E., Bogacka, I., Smith, S. R., Hulver, M., Ravussin, E. (2005) Glucose tolerance and skeletal muscle gene expression in response to alternate day fasting. Obes. Res. 13,574-581[Medline]
  70. Nisoli, E., Clementi, E., Paolucci, C., Cozzi, V., Tonello, C., Sciorinati, C., Bracale, R., Valerio, A., Francolini, M., Moncada, S., et al (2003) Mitochondrial biogenesis in mammals: the role of endogenous nitric oxide. Science 299,896-899[Abstract/Free Full Text]
  71. Nisoli, E., Tonello, C., Cardile, A., Cozzi, V., Bracale, R., Tedesco, L., Falcone, S., Valerio, A., Cantoni, O., Clementi, E., et al (2005) Calorie restriction promotes mitochondrial biogenesis by inducing the expression of eNOS. Science 310,314-317[Abstract/Free Full Text]
  72. Sauve, A. A., Wolberger, C., Schramm, V. L., Boeke, J. D. (2006) The biochemistry of sirtuins. Annu. Rev. Biochem. 75,435-465[CrossRef][Medline]
  73. Gerhart-Hines, Z., Rodgers, J. T., Bare, O., Lerin, C., Kim, S. H., Mostoslavsky, R., Alt, F. W., Wu, Z., Puigserver, P. (2007) Metabolic control of muscle mitochondrial function and fatty acid oxidation through SIRT1/PGC-1alpha. EMBO J. 26,1913-1923[CrossRef][Medline]
  74. Lagouge, M., Argmann, C., Gerhart-Hines, Z., Meziane, H., Lerin, C., Daussin, F., Messadeq, N., Milne, J., Lambert, P., Elliott, P., et al (2006) Resveratrol improves mitochondrial function and protects against metabolic disease by activating SIRT1 and PGC-1{alpha}. Cell 127,1109-1122[CrossRef][Medline]
  75. Shi, T., Wang, F., Stieren, E., Tong, Q. (2005) SIRT3, a mitochondrial sirtuin deacetylase, regulates mitochondrial function and thermogenesis in brown adipocytes. J. Biol. Chem. 14,13560-13567
  76. Mornagui, B., Grissa, A., Duvareille, M., Gharib, C., Kamoun, A., El-Fazaa, S., Gharbi, N. (2006) Vasopressin and nitric oxide synthesis after three days of water or food deprivation. Acta Biol. Hung. 57,1-11[CrossRef][Medline]
  77. Chen, Z. P., McConell, G. K., Michell, B. J., Snow, R. J., Canny, B. J., Kemp, B. E. (2000) AMPK signaling in contracting human skeletal muscle. Acetyl-CoA carboxylase and NO synthase phosphorylation. Am. J. Physiol. Endocrinol. Metab. 279,E1202-E1206[Abstract/Free Full Text]
  78. Stephens, T. J., Canny, B. J., Snow, R. J., McConell, G. K. (2004) 5'-aminoimidazole-4-carboxyamide-ribonucleoside-activated glucose transport is not prevented by nitric oxide synthase inhibition in rat isolated skeletal muscle. Clin. Exp. Pharmacol. Physiol. 31,419-423[CrossRef][Medline]
  79. Zou, M. H., Hou, X. Y., Shi, M., Kirkpatrick, S., Liu, F., Goldman, M. H., Cohen, R. A. (2003) Activation of 5'AMP-activated kinase is mediated through c-Scr and phosphoinositide 3-kinase activity during hypoxia-reoxygenation of bovine aortic endothelial cells. Role of peroxynitrite. J. Biol. Chem. 279,20767-20774[CrossRef]
  80. Shearer, J., Fueger, P. T., Vorndick, B., Bracy, D. P., Rottman, J. N., Clanton, J. A., Wasserman, D. H. (2004) AMP kinase-induced skeletal muscle glucose but not long-chain fatty acid uptake is dependent on nitric oxide. Diabetes 53,1429-1435[Abstract/Free Full Text]
  81. Jungas, R. L., Halperin, M. L., Brosnan, J. T. (1992) Quantitative analysis of amino acid oxidation and related gluconeogenesis in humans. Physiol. Rev. 72,419-448[Abstract/Free Full Text]
  82. Rasmussen, B. B., Wolfe, R. R. (1999) Regulation of fatty acid oxidation in skeletal muscle. Annu. Rev. Nutr. 19,463-484[CrossRef][Medline]
  83. Ruderman, N. B., Saha, A. K., Kragen, E. W. (2003) Minireview: Malonyl CoA, AMP-activated protein kinase, and adiposity. Endocrinology 144,5166-5177[Abstract/Free Full Text]
  84. Bonen, A., Luiken, J. J., Arumugam, Y., Glatz, J. F., Tandon, N. (2000) Acute regulation of fatty acid uptake involves the cellular redistribution of fatty acid translocase. J. Biol. Chem. 275,14501-14508[Abstract/Free Full Text]
  85. Luiken, J. J., Coort, S. L. M., Willems, J., Coumans, W. A., Bonen, A., van der Vusse, G. J., Glatz, J. F. (2003) Contraction-induced fatty acid translocase/CD36 translocation in rat cardiac myocytes is mediated through AMP-activated protein kinase signaling. Diabetes 52,1627-1634[Abstract/Free Full Text]
  86. Campbell, S. A., Tandon, N. A., Woldegiorgis, G., Luiken, J. J. F. P., Glatz, J. F. C., Bonen, A. (2004) A novel function for fatty acid translocase (FAT)/CD36. J. Biol. Chem. 279,36235-36241[Abstract/Free Full Text]
  87. Bezaire, V., Spriet, L. L., Campbell, S., Sabet, N., Gerrits, M., Bonen, A., Harper, M. E. (2005) Constitutive UCP3 overexpression at physiological levels increases mouse skeletal muscle capacity for fatty acid transport and oxidation. FASEB J. 19,977-979[Abstract/Free Full Text]
  88. MacLellan, J. D., Gerrits, M. F., Gowing, A., Smith, P. J. S., Wheeler, M. B., Harper, M. E. (2005) Physiological increases in uncoupling protein 3 augment fatty acid oxidation and decrease reactive oxygen species production without uncoupling respiration in muscle cells. Diabetes 54,2343-2350[Abstract/Free Full Text]
  89. Goglia, F., Skulachev, V. P. (2003) A function for novel uncoupling proteins: antioxidant defense of mitochondrial matrix by translocating fatty acid peroxides from the inner to the outer membrane leaflet. FASEB J. 17,1585-1591[Abstract/Free Full Text]
  90. Solanes, G., Pedraza, N., Iglesias, R., Giralt, M., Villaroya, F. (2000) The human uncoupling protein-3 gene promoter requires MyoD and is induced by retinoic acid in muscle cells. FASEB J. 14,2141-2143[Free Full Text]
  91. Solanes, G., Pedraza, N., Iglesias, R., Giralt, M., Villarroya, F. (2003) Functional relationship between MyoD and peroxisome proliferator-activated receptor-dependent regulatory pathways in the control of the human uncoupling protein-3 gene transcription. Mol. Endocrinol. 217,1944-1958
  92. Kong, Y., Flick, M. J., Kudla, A. J., Konieczny, S. F. (1997) Muscle LIM protein promotes myogenesis by enhancing the activity of MyoD. Mol. Cell. Biol. 17,4750-4769[Abstract]
  93. Khurana, T., Khurana, B., Noegel, A. A. (2002) LIM proteins: association with the actin cytoskeleton. Protoplasma 219,1-12[CrossRef][Medline]
  94. Schrauwen, P., Hardie, D. G., Boorda, B., Clapham, J. C., Abuin, A., Thomason-Hughes, M., Green, K., Frederik, P. M., Hesselink, M. K. (2004) Improved glucose homeostasis in mice overexpressing human UCP3: a role for AMP kinase?. Int. J. Obes. 28,824-828[CrossRef][Medline]
  95. Freyssenet, D. (2007) Energy sensing and regulation of gene expression in skeletal muscle. J. Appl. Physiol. 102,529-540[Abstract/Free Full Text]
  96. Randle, P. J., Garland, P. B., Hales, C. N., Newsholme, E. A. (1963) The glucose fatty-acid cycle. Its role in insulin sensitivity and the metabolic disturbances of diabetes mellitus. Lancet 1,785-789[Medline]
  97. Petersen, K. F., Shulman, G. I. (2006) Etiology of insulin resistance. Am. J. Med. 119,10S-16S[CrossRef][Medline]



This article has been cited by other articles:


Home page
 EndocrinologyHome page
S.-i. Oka, E. Yoshihara, A. Bizen-Abe, W. Liu, M. Watanabe, J. Yodoi, and H. Masutani
Thioredoxin Binding Protein-2/Thioredoxin-Interacting Protein Is a Critical Regulator of Insulin Secretion and Peroxisome Proliferator-Activated Receptor Function
Endocrinology, March 1, 2009; 150(3): 1225 - 1234.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
P. de Lange, R. Senese, F. Cioffi, M. Moreno, A. Lombardi, E. Silvestri, F. Goglia, and A. Lanni
Rapid Activation by 3,5,3'-L-Triiodothyronine of Adenosine 5'-Monophosphate-Activated Protein Kinase/Acetyl-Coenzyme A Carboxylase and Akt/Protein Kinase B Signaling Pathways: Relation to Changes in Fuel Metabolism and Myosin Heavy-Chain Protein Content in Rat Gastrocnemius Muscle in Vivo
Endocrinology, December 1, 2008; 149(12): 6462 - 6470.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Endocrinol. Metab.Home page
C. Roy, S. Paglialunga, A. Fisette, P. Schrauwen, E. Moonen-Kornips, J. St-Onge, M. K. Hesselink, D. Richard, D. R. Joanisse, and K. Cianflone
Shift in metabolic fuel in acylation-stimulating protein-deficient mice following a high-fat diet
Am J Physiol Endocrinol Metab, June 1, 2008; 294(6): E1051 - E1059.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
Z. Nahle, M. Hsieh, T. Pietka, C. T. Coburn, P. A. Grimaldi, M. Q. Zhang, D. Das, and N. A. Abumrad
CD36-dependent Regulation of Muscle FoxO1 and PDK4 in the PPAR{delta}/{beta}-mediated Adaptation to Metabolic Stress
J. Biol. Chem., May 23, 2008; 283(21): 14317 - 14326.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
fj.07-8527revv1
fj.07-8527revv2
21/13/3431    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pieter de Lange
Right arrow Articles by Lanni, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pieter de Lange,
Right arrow Articles by Lanni, A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS